






















































































• A v-* \v 

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IWY? 

ys 


an evaluation of 
CHESTER RIl'ER 
OYSTER /MORTALITY 


JAN 0 4" 1964 

CLEARED MCL 





/M/4RYL/4ND DER4RT/HENT OF N/4TUR/4L RESOUCES 
















































AN EVALUATION OF CHESTER RIVER OYSTER MORTALITY 


Joseph J. Cooney, F, Douglas Martin, and W. H. Roosenberg 
Chesapeake Biological Laboratory, University of Maryland 

Solomons, Maryland 20688 

and 


David H. Freeman 

Department of Chemistry and Chesapeake Biological Laboratory 

College Park, Maryland 20742 

and 

Charles R. Bostater, Jr. 

Project Manager, Maryland Water Resources Administration 

Annapolis, Maryland 21401 


Funded By 

Chesapeake Bay Program Grant #805976 

and 

Maryland Department of Natural Resources 

Lowell Bahner, Project Officer 
EPA Chesapeake Bay Program 
Gulf Breeze Environmental Research Laboratory 
Annapolis, Maryland 21401 


GULF BREEZE ENVIRONMENTAL RESEARCH LABORATORY 
OFFICE OF RESEARCH AND DEVELOPMENT 
U.S. ENVIRONMENTAL PROTECTION AGENCY 
GULF BREEZE, FLORIDA 32561 




DISCLAIMER 


This report has been reviewed by the Chesapeake Bay Program, U.S. 
Environmental Protection Agency, and approved for publication. Approval 
does not signify that the contents necessarily reflect the views and 
policies of the U. S. Environmental Protection Agency or the Maryland 
Department of Natural Resources, nor does mention of trade names or 
commercial products constitute endorsement or recommendation for use. 



.7 



PREFACE 


\ 


The Maryland Water Resources Administration received a 
grant from the U.S. EPA Chesapeake Bay Program to conduct a 
study concerning oyster mortality in the Chester River, a sub- 
estuary of the Chesapeake Bay. The State of Maryland contracted 
with the University of Maryland to perform a three—part study to 
evaluate the reported oyster mortality. Due to the fact that 
this was a retrospective study, the evaluation of past large 
instananeous inputs of chemicals and the resulting potential 
impact was not made. The results of the University of Maryland's 
research follows. 

The results of the three studies involving aquatic bioassays 
and arlalysis of phthalate esters, tin and organotin compounds in 
various environmental media did not show conclusive cause-and- 
effect relationships concerning the oyster mortality. 

The bioassay study showed no significant point source or 
ambient acute aquatic toxicity to organisms tested; however, 
chronic stress was indicated by growth reduction of oysters 
during the study period and by occasional low levels of dis¬ 
solved oxygen in the lower estuary. The point source bioassay 
observations showed mortality of organisms as well as controls; 
however, the controls and point source tests were not signifi¬ 
cantly different. 

The analysis of phthalate esters in alluvial sediments 
showed a decreasing trend downstream from a point source. A 
pond receiving an industrial discharge showed extremely high 
concentrations of phthalate esters. The levels found in the 
vicinity of the pond warrant further consideration. Levels of 
phthalate esters in the estuarine sediments are low. The tox¬ 
icity of the levels found is unknown, especially to oysters. 

A review of current literature shows no documentation of the 
toxicity and dynamics of phthalates to exposed oysters. The 
oyster concentration data in this report are the first published 
data available for di (2-ethylhexyl) phthalate (DEHP) in the 
American oyster. Although the toxicity of phthalates to the 
oyster would be expected to be low, because of the economic 
importance of the oyster, further laboratory bioassays are 
warranted to determine acute-and chronic-effect concentration 
levels. 

Other modeling efforts conducted by state and Federal 
- workers have estimated that the major transport pathway of 


IV 


phthalate esters is in the dissolved fraction in the water 
column. It is not known at this time if the water phase trans¬ 
port of phthalates out of the system and other environmental 
degradation processes are great enough to preclude further 
buildup of DEIIP in the estuarine sediments. Based on the levels 
of DEHP found in the pond receiving an effluent, further accumu¬ 
lation of phthalates in the estuarine sediments may be possible. 
Additional modeling could address these issues if sediment 
deposition and resuspension calculations were included in a 
modeling frameworks. 

Tin, microorganisms resistant to tin, and microorganisms 
capable of transforming inorganic tin to organotin(s) were 
present at the sites sampled. However, the analysis of tin 
and organotin compounds in Chester River media did not result 
in any data to support or preclude the possibility of oyster 
mortality due to organotin compounds. 

In summary, no significant mortality of oysters was 
observed during the course of this study. However, there are 
indications of chronic stress in the estuarine system based on 
the results of this study. Should mortality be observed again 
at any time in the future, it is recommended that oyster samples 
should be taken immediately, stored, and later analyzed for 
suspected xenobiotics. At the same time, water quality variables 
such as dissolved oxygen, salinity and other environmental 
variables should be observed as soon as possible after the 
mortality is reported. It should be noted that the greater the 
time period between when mortality of organisms is reported, and 
the analytical observations are made, the more difficult it will 
be to evaluate possible cause-and-effect relationships. 

In addition, the acute and chronic effects of phthalate 
esters to estuarine organisms is relatively unknown. Additional 
monitoring of phthalates in this River is recommended as well 
as laboratory aquatic bioassay tests. Monitoring of oyster 
growth and mortality, dissolved oxygen and salinity over a 
several year period is required to determine if the observed 
high mortality has subsided. Such monitoring would help to 
identify causes of chronic stress indicated by reduced growth 
rate of the oysters. 


v 


ABSTRACT 


EVALUATION OF CHESTER RIVER MORTALITY BIOTOXICITY 

i, 

Three studies were performed to determine whether the recent 
dieoffs of oysters in the Chester River can be correlated with 
point sources of toxic substances. To this end two kinds of ex¬ 
periments were performed: long-term experiments with oysters 
placed in the Chester River; and 96-hour acute toxicity experi¬ 
ments with golden shiners, Notemigo nus chrysoleucas, and cravfish. 
Procambarus acutus acutus . - 

In the long-term studies, 10 stations were established. 

Three stations were below the areas of known oysterkills, four 
were within the areas of recent oysterkills and three were in 
areas where the kills occurred in 1974 and 1975. One or two trays 
with 96 oyster specimens were placed at each station. Ten or 
twelve oysters were removed at nine intervals during 4 months. 
These were scored for condition. No significant mortality occur¬ 
red during this period, but during July and August 1978 the five 
stations most upriver had dieoff of the fouling organisms and 
reduced growth rates of the oysters. 

The 96—hour acute toxicity studies were performed by placing 
cages of golden shiners and crayfish in streams receiving ef¬ 
fluents from the Campbell's Soup plant, the Tenneco, Inc. plant 
and the sewage treatment plant for the city of Chestertown. No 
significant mortality occurred. 

No point sources of toxicants were located, but since no 
significant mortality occurred during the study these results are 
not conclusive. 

PHTHALATE ESTERS AND RELATED CHEMICALS IN THE CHESTER RIVER BASIN 

The Tenneco factory on Morgan Creek is permitted to discharge 
up to 2830 kg of organic extractables per year. These waste 
chemicals empty into the adjacent Tenneco Pond with a residence 
time of ca. 10^ days. The pond discharge flows through Morgan 
Creek into the Chester River. The residence time in the river is 
a much shorter 130 days. 

Model concepts based on available data allow plausible calcu¬ 
lations of discharge organics in the contiguous downstream sedi¬ 
ments. The possibilities range from 0.04 ppm based on simple 


vi 







partition to 5 ppm for a "hot spot" model based on previous dye 
discharge experiments. 

New chemical methods based on gas chromatography/mass spec¬ 
trometry analysis of DBP (dibutylphthalate) and DEHP (di(2- 
ethylhexyl)phthalate) were developed. The relative standard de¬ 
viation was demonstrated at + 20 percent of the 0.1 ppm level in 
sediment. Accuracy is more difficult to specify, but this may 
be judged on the basis of split samples measured by an independ¬ 
ent laboratory (0.3 ppm) and our own data (0.1 ppm). 

MICROBIAL TRANSFORMATION OF TIN 

This work was undertaken to determine if tin or products re¬ 
sulting from the biotransformation of tin may contribute to oys¬ 
ter mortality in the Chester River, Maryland. 

Data were collected at two sites in the River: Spaniard 
Bar, which suffered extensive oyster mortality, and Buoy Rock, 
which did not exhibit extensive oyster mortality. Three sites 
associated with potential sources of tin in the River were also 
studied: the Tenneco plant and the Campbell's Soup plant, both 

near Chestertown, Maryland, and the Chestertown sewage treatment 
plant. For comparison, some samples were taken in Baltimore 
Harbor, a site known to be polluted with heavy metals, and in 
Tangier Sound near Tilghman Island, a site regarded as relatively 
free of pollution. 

Water and sediment samples were examined for total viable 
counts of microorganisms, for counts of microorganisms resistant 
to inorganic tin, and for counts of microorganisms resistant to 
organic tin. Sediment from each site was used as inoculum for 
cultures to determine if microorganisms at the site could trans¬ 
form inorganic tin to volatile (organic) tin compound(s). Water 
and sediment were assayed for tin content. 

Among physiochemical parameters measured onsite, only low 
dissolved oxygen is a potential contributor to oyster mortality. 
Microorganisms resistant to inorganic tin were detected in all 
samples and most samples contained microorganisms resistant to 
organotin, although organotin was more toxic than inorganotin to 
the microbial flora. Microorganisms capable of converting in¬ 
organic tin to volatile tin compound(s) were present at every 
site. Comparison of tin concentrations at the several sites 
showed that it is not possible to attribute the oyster kill 
solely to tin, although interaction with other stress factors is 
possible. 

This work was submitted in fulfillment of contract # 
R805976010 by the University of Maryland, Chesapeake Biological 
Laboratory under contract to the Maryland Water Resources Admin¬ 
istration and sponsored by the U.S. Environmental Protection 


Vll 


Agency. This report covers the period January 1, 1978 to July 
23, 1979, and work was completed as of August 6, 1979. 






































































» 



















• • • 




Vlll 







CONTENTS 



Foreword.iii 

Preface.iv 

Abstract.vi 

Figures.xi 

Tables.xii 

Acknowledgement . xiv 


1. Introduction . 1 

Evaluation of biotoxicity . 1 

Phthalate esters and related chemicals . 1 

Microbial transformation of tin . 2 

2. Conclusions . :.'. 5 

Evaluation of biotoxicity . 5 

Phthalate esters and related chemicals . 5 

Microbial transformation of tin . 6 

3. Recommendations. 8 

Evaluation of biotoxicity . 8 

Phthalate esters and related chemicals . 8 

Microbial transformation of tin . 9 

4. Evaluation of Chester River Oyster Mortality 

Biotoxicity.10 

Methods and Materials . 10 

Procedures.17 

Results.24 

Discussion.36 


IX 

























39 


5. Phthalate Esters and Related Chemicals in 

the Chester River . 

Introduction . 39 

Distribution Patterns . 43 

Problem Statement . 43 

Experimental Procedures . 44 

Results and Discussion . 56 

6. Microbial Transformation of Tin. 87 

Experimental Procedures . 87 

Results and Discussion . 91 

Appendices 

A. Chester River Oyster Mortality . 103 

B. Account of Interlaboratory Tests on Split Samples. 112 

C. Methods for Water Quality Measurements in 

Tin Study.114 


x 













FIGURES 


Number Page 

1 Lower Chester River showing oyster tray stations 

1 thru 6. 12 

2 Lower Chester River showing oyster tray stations 

6 thru 10. 13 

3 Mortality of golden shiners, by station, over the 

96-hour observation period . 31 

4 Excess mortality of golden shiners, by station, 

during 96 hours of observation . 33 

5 Map of Chester River illustrating sample sites ... 40 

6 Downstream progession of oyster mortality . 41 

7 GCMS of Chester River mouth sediment extract .... 46 

8 Liquid chromatography of Tenneco Products designated 

by Tenneco as Alkyl Phthalates, or mixtures 
(6-10P and 7-11P). 57 

9 Liquid chromatogram of DOP, DIDP, and DTDP 

(synthetic mixture of Tenneco samples) . 58 

10 Liquid chromatogram of Tenneco pond water . 59 

11 Liquid chromatogram of Tenneco pond sediment .... 61 

12 Liquid chromatogram of Morgan Creek sediment from 

Frye Farm a few miles downstream from Tenneco ... 63 

13 Comparison liquid chromatogram of humic acid .... 63 

14 Comparison of GCMS-SIM chromatograms . 70 

15 Oyster tissue extract (spiked with 20 ppm DEP, 

DBP, and DEHP) . 78 

16 Oyster tissue extract . 79 


xi 















TABLES 


'c 

Number Page 

1 Physical Parameters Measured at Chester River 

Oyster Study Sites . 14 

2 Hydrographic Observations for the Possible Point 

Source Studies . 22 

3 Mortality of Oysters During This Study . 24 

4 Comparison of Mean New Growth in Oysters Planted 

in Chester River. 25 

5 Comparison of Mean Observed Condition Scores in 

Oysters Planted in Chester River . 27 

6 Associated Organisms by Station . 28 

7 Relationship of Nonsurvival to Autopsy Data .... 34 

8 Comparisons between Stations for Characters Related 

to Nonsurvival. 35 

9 Field Sample Handling . 52 

10 Methodology. 53 

11 Effect of Sonication on DBP Extraction by CH-Cl^ 

from Sample "R" .7 . . 62 

12 Comparison of Ultrasonic Extraction Efficiencies 

of Varied Solvents Measured on Working Standard . 64 

13 DEHP Recovery Measurements Using Dichloromethane . . 65 

14 Comparison of Methods for Extraction of Phthalate 

Esters from Chester River, Mouth Sediment, 

Test 1. 68 

15 Test of Varied Extraction and Re-extraction 

Procedures. 68 


Xll 
















Number Page 

16 Interlaboratory Comparison of DEHP Measurements 

of Split Samples.71 

17 Determination of Sediment Composition . 73 

18 Concentration of Phthalates in Oysters and 

Related Sediments . 80 

19 Determination of a Concentration of Tin Which Would 

Select for Tin-Resistant Microorganisms . 89 

20 Physical Data for April 1978 Cruise/Excursion .... 92 

21 Physical Data for July 1979 Cruise/Excursion . 92 

22 Viable Counts of Bacteria from April 1978 Cruise ... 93 

23 Viable Counts of Bacteria from July 1979 Cruise ... 94 

24 Production of Volatile Tin in Media Inoculated 

with Sediment.96 

25 Tin in Water and Sediment Samples.97 


xm 











ACKNOWLEDGEMENTS 


The Evaluation of Chester River Oyster Mortality Biotoxicity 
study was performed by F. Douglas Martin and Willem H. Roosenburg 
of the University of Maryland, Center for Environmental and Estu¬ 
arine Studies, Chesapeake Biological Laboratory, Solomons, Mary¬ 
land, 20688. They wish to thank Dr. George Krantz of UM/CEES 
Horn Point Laboratories who advised on appropriate location of 
stations; Robert Miller, Daniel Carver, Sharon Ambrose, Dorothy 
Cutshaw, Wayland Owens, Rev. Mark Odell and others who assisted 
in field work; and Captain William Keefe and several Kent County 
residents for rescue services. 

David H. Freeman, Department of Chemistry, University of 
Maryland, College Park, Maryland, 20742, authored the study on 
Phthalate Esters and Related Chemicals in the Chester River 
Basin. The experimental work in this study was carried out by 
James C. Peterson and Sally A. Gingras. The work depended 
critically upon ultraclean glassware and John Trembly cooperated 
by facilitating routine use of the Chemistry Department's an¬ 
nealing ovens for this purpose. Mr. Arden Fox of Tenneco was 
helpful in providing details and insights to the history and 
present operations of the Tenneco plant and its bacterial waste 
processing facility. Captain O'Berry and the crew of the 
"Aquarius" ably assisted the collection of the Chester River 
water and sediment samples. The research sailing vessel "Huckle¬ 
berry Friend" and the facilities of the Podickory Sailing Assoc¬ 
iation were used to gather the reference samples from the mouth 
of the Chester River. Special thanks go to Dr. William Budde 
who generously provided cooperative measurements on split sedi¬ 
ment samples. Finally, we happily acknowledge the value of 
Professor Ron Hites' intensive short course on environmental 
applications of GCMS. Drs. Nelson Frew, Robert Gagosian and 
John Farrington at Woods Hole Oceanographic Institution provided 
helpful consultation during the course of this work. 

Microbial Transformation of Tin was under the direction of 
Joseph Cooney, Chesapeake Biological Laboratory, Center for 
Environment and Estuarine Studies, University of Maryland, 
Solomons, Maryland, 20688. Experimental work was conducted by 
L. E. Hallas, with the assistance of Marthe Cole-Jones and 
Terri Ekelund. 


xiv 


SECTION 1 


INTRODUCTION 


EVALUATION OF BIOTOXICITY 

The Chester River is a tributary of the Chesapeake Bay, 
northeast of Kent Narrows. Historically, this estuarine part of 
the river has been rich in natural oyster bars; but, in the 
winter of 1974, a heavy mortality started upriver and moved over 
a period of years to bars in the lower reaches. The symptoms of 
this oyster mortality could not be attributed to any climatic or 
other natural phenomena that cause occasional oyster mortalities. 
A suspicion became plausible that something highly toxic to 
oysters had entered the upper river and had gradually worked its 
way downstream. A review of possible point sources of waste dis¬ 
charge into the Chester River system revealed the presence of the 
Chestertown sewage plant on Radcliffe Creek, the Campbell's Soup 
factory on Morgan Creek and the Tenneco plant that dumps its ef¬ 
fluent into a pond which eventually empties into Morgan Creek via 
a small creek. The investigations reported here were aimed at 
locating point sources of toxic materials, and stations were 
chosen to maximize the ability to identify the roles of these 
potential sources. 

PHTHALATE ESTERS AND RELATED CHEMICALS 

The State of Maryland has issued a permit to Tenneco, In¬ 
corporated, in Chestertown, Maryland, to discharge 10 ppm of 
total organic extractables into Morgan Creek which empties into 
the Chester River and, then, into Chesapeake Bay. This permit 
would appear to be reasonably conservative unless it conceals the 
basis for long-range harm. Such threatening possibilities do 
exist and will be considered in the present studies. The major 
problem is related to the question of whether the discharge is 
free to dilute itself in some innocuous way, or whether, on the 
contrary, the dilution processes are blocked and the basis for a 
toxic accumulation can be shown. In the latter case, the permit 
would have to be viewed as nonconservative and the ecological 
threats would have to be carefully considered and perhaps further 
constrained. 


1 


The present study includes an initial investigation of 
various models for the distribution of the chemicals discharged 
from the Tenneco site. Various options are considered between 
the two extremes—that the organics are fully trapped at the 
Tenneco site, or that all the organics are distributed uniformly 
into the Chester River sediment beds. 

The chemical analysis of sediment and oyster tissue for alkyl 
phthalates is prone to a myriad of error sources due to ubiqui¬ 
tous presence of these plasticizer compounds in the laboratory. 

As a result, there is a need to develop new technology that would 
scrupulously suppress the opportunity for contamination. The 
development succeeded because it relies on a well-established 
statistical maxim--that any risk if repeated often enough must 
lead to disaster. Since chemical methodology consists of a 
series of steps, each with an assigned risk, the approach was to 
develop a procedure that was designed to approach zero reliance 
on chemical manipulations. 

Quality assurance was not a part of the original work plan. 
However, independent laboratory tests of split samples showed 
that the developed chemical methodology was indeed adequate for 
the purposes at hand. 

The underlying goals, then, were to develop and test the new 
methodology, to explore the possible causal link of industrially 
discharged alkyl phthalates to the past oyster mortality, and to 
determine whether the present discharge constitutes an ecological 
threat. 

MICROBIAL TRANSFORMATION OF TIN 

Organotin compounds were first synthesized about 1850 (Van 
der Kerk 1976) , and they were first used as agents to control 
biological activity around 1930, when they were used as moth¬ 
proofing agents (Luijten 1972) . Shortly thereafter, organotins 
were used as stabilizers for vinyl resins, which continues to be 
a major application (Subramanian 1978). In the last 10 years, 
use of tin by industrial societies has more than doubled. Or¬ 
ganotin compounds are used widely to control a variety of plants, 
animals, and microorganisms (Deschiens and Floch 1962, Daum 1965, 
Holden 1972, Luijten 1972, van der Kerk 1976). All such organo¬ 
tin compounds are toxic, but the effect varies with the organic 
group(s) present (Thayer 1974). In general, triorganotin com¬ 
pounds are more toxic than di- or tetraorgano compounds. Dior- 
ganotins behave like organomercurials, reacting with sulfhydryl 
groups to inactivate enzymes. Trialklyl tin compounds interfere 
with oxidative phosphorylation and with photosynthetic phos¬ 
phorylation (Thayer 1974). Methyl tin compounds are poisonous 
to the central nervous systems of higher organisms (Ridley, 
Dizikes, and Wood 1977). Effects can be species-specific. For 


2 


* 


example, at concentrations too low (15-30 ppb) to affect some 
fish, triphenyltin acetate and several tributyltin compounds af¬ 
fect snails, zooplankton, and small fish, while warm-blooded 
species show no toxicity. In contrast, some soil microorganisms 
are not affected by concentrations of tributyltin oxide at con¬ 
centrations up to 100 ppm (Thayer 1974) . 

In comparison with other metals such as mercury, lead, and 
cadmium, relatively little is known about biological transforma¬ 
tions of tin. But sufficient information is available that a 
biological cycle has been proposed (Ridley et al. 1977). 



b 


(a) CH 3 SnX 3 + e-+CH 3 —»-(CH 3 ) 2 SnX 2 + X" 


In the diagram, X indicates a counteranion. The free radical 
SnX 3 can be methylated in a series of biological reactions (re¬ 
action a) involving methylocobalamins (e.g., vitamin-B]_ 2 ) to 
yield mono-, di-, tri-, and tetramethyl tin (Ridley et al. 1977). 
A Pseudomonas species which is purportedly capable of methylating 
tin has been isolated from Chesapeake Bay (Huey et al. 1974). In 
the presence of ionic mercury, trimethyltin can react to yield 
the highly toxic methylmercury (Huey et al. 1974). Methyltins 


3 












could be cleaved oxidatively (reaction b) by mixed function 
oxidases in the same way that trialkylyltin derivates are 
cleaved by liver microsomes (Kimmel, Fish, and Casida 1977). 
Alkyltins can also be degraded by photolysis (*) to yield free 
radicals (Lloyd and Rogers 1973). 

Thus, tin can be transformed biologically to toxic compounds 
and these toxic compounds can be transformed chemically to other 
toxic compounds. Since tin is used at the Tenneco plant near 
Chestertown, it is a potential source of compounds toxic to 
oysters. 

The objective of the present study was to determine if tin 
or products resulting from the microbial transformation of tin 
could contribute to oyster mortality in the Chester River. The 
investigation was not designed as a definitive study, but as a 
preliminary, screening study to determine whether further studies 
are warranted. 





4 


SECTION 2 


CONCLUSIONS 


EVALUATION OF BIOTOXICITY 

Since there was no significant mortality in our experimental 
oysters, there was no strong indication that the causative factor 
for oysterkills in the Chester River was in operation during our 
studies. 

Fouling organisms died off in July and August at five sta¬ 
tions upriver from Corsica Neck. Since this accompanied dieoff 
of oysters in previous years, the phenomenon which is responsible 
for oyster dieoff might have occurred but in a mild form. 

Despite reasonable growth in the oysters planted in Chester River, 
the growth rate during the period of fouling community dieoff was 
significantly lower than that of controls placed in the Patuxent 
River. The phenomenon this year may have been so mild that the 
main effect in the oysters was a reduction in new growth. 

The beginning of dieoff of the fouling community correlated 
with a fishkill. The fishkill originated well above Morgan Creek. 
The two phenomena may be unrelated as the fishkill may be bacte¬ 
rial in origin and species-specific since only carp and catfishes 
were noted dying. 

Experiments with fish and crayfish in Radcliff Creek and 
Morgan Creek do not find any indication that either creek con¬ 
tains the sole source of the cause of oyster mortality within its 
drainage; however, the lack of significant mortality during the 
study period makes these results inconclusive. 

PHTHALATE ESTERS AND RELATED CHEMICALS 

The Chester River sediments taken from the vicinity of the 
oyster mortality zone, as well as farther downstream, show no 
evidence that Tenneco discharges are causally linked to the past 
oyster mortality. Oysters are now grown with apparent health in 
regions where the alkyl phthalates should be similar in concen¬ 
tration to those presently measured in the vicinity of the oyster 
mortality. Since the concentration history has not been measured, 
the study cannot rule out the possibility of a past causal re¬ 
lationship. 


5 


A massive buildup of alkyl phthalates manufactured by Tenneco 
was found in Tenneco Pond immediately adjacent to the factory 
waste water discharge plant. The levels show clearly that the 
pond sediments are serving as a sink, i.e., as a secondary waste 
treatment facility. 

Experimental measurements of Chester River sediments show no 
significant differences between the mortality zone and the Ches¬ 
ter River mouth where reasonably healthy oyster growth persists. 
The results for DBP and DEHP are very nearly identical in these 
regions in the Chester River: 0.02-0.85 ppm with average values 
of DBP (0.5 ppm) and DEHP (0.05 ppm). The DEHP/DBP ratio is 
0.1 + 0.07. 

The Tenneco Pond results are quite different: DEHP (1.5 x 
10^ ppm) and DBP (0.2 ppm). Tenneco has rarely made DBP. The 
difference in the DEHP/DBP ratio alone suggests that the alkyl 
phthalates in the Chester River may originate from Tenneco as 
well as other possible sources. Moreover, the estimated possible 
accumulation in Tenneco Pond—10^ kg of alkyl phthalates—sug¬ 
gests that the pond functions in part as a waste treatment 
facility. 

The greatest threat seems to be that the Tenneco Pond may be 
nearing the saturation state that it must reach eventually. In 
that case, one can confidently forecast a serious accumulation 
of the alkyl phthalates that will in time spread out into the 
Morgan Creek area. 

MICROBIAL TRANSFORMATION OF TIN 

Examination of physiochemical data (pH, temperature, 
salinity, dissolved oxygen) from four estuarine and three fresh¬ 
water sites shows only low dissolved oxygen near the bottom as a 
potential contributor to oyster mortality in the Chester River. 

All sediment and water samples examined contain microorga¬ 
nisms resistant to inorganic tin; resistant organisms comprised 
as much as 55 percent of the total aerobic, heterotrophic popu¬ 
lation detected. Most of the water and sediment samples con¬ 
tained organisms resistant to the organotin compound, 
dimethyltin chloride; such organisms comprised as much as 17 
percent of the total aerobic, heterotrophic population detected. 

Microbial populations are more sensitive to organotin than 
to inorganic tin. 

Microorganisms capable of converting inorganic tin to 
volatile tin compound(s) are widely distributed in the Chesa¬ 
peake ecosystem. 


6 


All sediments associated with the Chester River--including 
sediments from the Tenneco plant, the Campbell plant, and the 
Chestertown sewage treatment plant—contained more tin than sedi¬ 
ment from a site near Tilghman Island and less tin than a site in 
Baltimore Harbor. Spaniard Bar, which suffered an oysterkill, 
did not yield significantly more than Buoy Rock, which did suffer 
such a kill. Thus, it is not possible to attribute the oyster- 
kill in the Chester River solely to pollution by tin, although 
interaction with other stress factors is possible. In addition 
to sediment, water in the Chester River and water entering the 
Chester River from the Tenneco plant, from the Campbell plant, 
and from the Chestertown sewage treatment plant sometimes contain 
significant quantities of tin. 

Significant progress has been made toward developing a method 
for separation and qualitative and quantitative measurement of 
organotin species in environmental samples and in microbial cul¬ 
tures . 


7 


SECTION 3 


RECOMMENDATIONS 


EVALUATION OF BIOTOXICITY 

Possible links between the dieoff of associated organisms and 
previous oysterkills should be further investigated. In addition 
to monitoring fouling communities of oysters, transite sheets 
should be placed on or near oyster bars and be sampled periodi¬ 
cally. These sheets make quantification of the fouling organisms 
simpler and quicker when the effects are not gross. 

The slower oyster growth in Chester River may be related to 
the oysterkill causal factor(s). Year-round sampling of oyster 
bars in the Chester River to monitor new growth is recommended. 

An area with a similar salinity regime and without such oyster- 
kill phenomenon should be sampled simultaneously as a control. 

The studies recommended above could be performed concurrently. 

A study continuously monitoring dissolved oxygen during 
periods when oysterkill may occur is recommended. 

PHTHALATE ESTERS AND RELATED CHEMICALS 

The levels of alkyl phthalates observed in the Chester River 
do not appear to present an immediate threat. However, the oyster 
lives a perilous existence. It is quite possible, and perhaps 
likely, that the alkyl phthalate levels are changing rapidly 
enough in the Chester River and the greater Chesapeake Bay to 
constitute a serious threat at some future time. 

It is recommended that the rate of buildup be monitored 
through piston cores taken in well-stratified sediments. The 
goal should be to forecast the date when the extrapolated levels 
will be reached where healthy oyster production will be prevented 
by the presence of these ubiquitous chemicals. 

It is recommended that the Morgan Creek sediments be surveyed 
for evidence of alkyl phthalate accumulation. The creek serves 
as a conduit between the Tenneco pond and contiguous farmland 
using the discharged water for irrigation purposes. The Tenneco 
Pond sediments are likely to be weakening in their role as a sink 
for these chemicals. Eventually, their sorptive capacity is 


8 


likely to become exhausted. 

It is recommended that Tenneco Pond be surveyed to establish 
the total phthalate plasticizer content. At the same time, co¬ 
operative research by Tenneco should be elicited to see if a more 
effective use of secondary waste water treatment by certain soils 
could be established in order to furnish long-range protection to 
the downstream ecology. 

MICROBIAL TRANSFORMATION OF TIN 

Tin should not be considered as the sole source of the exten¬ 
sive oyster mortality observed in portions of the Chester River. 

Tin should not be excluded as a partial cause of oyster mor¬ 
tality in the Chester River, particularly when coupled with other 
pollutants and with low dissolved oxygen. 

When tin enters an aquatic ecosystem, it should be assumed 
that microorganisms are present which can convert it to volatile 
tin compounds. 

Studies should be undertaken, using recently developed meth¬ 
odology, to determine if oysters bioaccumulate tin and if the 
oyster's gut flora can produce significant quantities of volatile 
tin compounds. 


9 


SECTION 4 


EVALUATION OF CHESTER RIVER OYSTER MORTALITY BIOTOXICITY 

METHODS AND MATERIALS 
Possible Sources of Toxicants 


The Chester River has a typical estuarine portion with widely 
varying temperature and salinity regimes. All of the possible 
sources of toxicants that were examined have their waste mate¬ 
rials entering the river in its tidal portion. These possible 
sources dump materials of widely differing nature. The Chester- 
town sewage plant discharges chlorinated and treated domestic 
sewage. The Campbell's Soup factory sprays its effluent, which 
are wastewaters from preparing chicken, over percolation fields 
and each leached water discharge is chlorinated before it enters 
into Morgan Creek. Tenneco does not discharge any effluent that 
can be made more acceptable by chlorination, as its effluent is 
resultant from manufacture of plasticizers. Instead of chlori¬ 
nating, the factory discharges into a leaching pond at one end 
with a stand pipe overflow on the far end. The volume of ef¬ 
fluent at the time of the study was a small volume compared to 
both the sewage plant and Campbell's Soup plant. 

The proposal called for two separate but related studies: 

(1) a field study on mortality and growth of oysters in the 
Chester River, and (2) a field study to detect possible toxic 
effects on test animals by the effluents of the three plants 
discharging into the Chester River basin. 

Oyster Studies-- 

The oyster portion of the study was conducted in the portion 
of the river from just below Chestertown to just above Kent 
Narrows, which is near the mouth. There were 10 stations in the 
Chester River from 15 May 1978 until 18 September 1978. Fifteen 
trays, each containing 96 oysters, were monitored for mortality. 
Qualitative information on oyster communities' associated orga¬ 
nisms were also recorded. 


10 



Stations--See Figures 1 and 2 for orientation of stations. 
Comments concerning these stations are given below. 


Station 1. 

North of Kent Island, a single-tray station. This 
tray was attached to parts of a sunken barge. At 
the first sampling visit the tray had been tampered 
with and when retrieved it came up upside down with 
the lid open. No oysters could be recovered and the 
station was abandoned. 

Station 2. 

South of Cedar Point. This was a two-tray station 
attached to clam buoy "SS." This station lasted 
from 15 May until 27 June but could not be found 
with the grapnel nor by diving on 12 July. 

Station 3. 

Off Tilghman Creek. This single-tray station was 

tied to a stake at 0.6 m depth. This station lasted 
the entire study period. 

Station 4. 

Off Piney Point. A double-tray station tied to clam 
buoy "C." This station served from 15 May - 4 

August but could not be found on the 23 August 
check and thereafter. 

Station 5. 

At Ringgold Point, a single-tray station, tied to a 
Coast Guard day marker by a nylon line to a ring on 
this structure. The line was cut by vandals before 
this station was visited on 27 June. The station 
was observed from 15 May - 14 June. 

Station 6. 

Off Corsica River. Thi^ station held two trays and 
was fastened to clam buoy "A." The tieline was at¬ 
tached to the buoy's anchor chain by means of a 
chain ring that dropped to the bottom. This sta¬ 
tion lasted the entire study period from 15 May to 

18 September. 

Station 7. 

Nichol's Point, a double-tray station. This station 
was tied to a ring on a day marker. It was in 
service for the entire 15 May - 18 September period. 

Station 8. 

Off Cliffs Point, a single-tray station attached to 
the remnants of a booby blind in about 1 m of water. 
The station lasted the entire study period. 

Station 9. 

Off the mouth of Shippen Creek. This single-tray 
station was attached to a privately owned pier and 


was not tampered with. 


Station 10. 

Newman's Wharf just off Deep point. This two-tray 
station was also suspended from a private pier and 
lasted the entire study period. 


11 



Figure 1. Lower Chester River showing oyster tray stations 
1 thru 6. 

\ 


12 














Ficfuire ^ * Lower Chester River shovTing oyster tray stations 
6 thru 10. 


13 















'*0 


Hydrographic observations of all stations are contained in 
Table 1. 


TABLE 1. PHYSICAL PARAMETERS MEASURED AT CHESTER 
RIVER OYSTER STUDY SITES 


Depth 


Surface/Bottom 


Secci 



(m) 

Salinity 

Temperature 

Dissolved O 2 

Depth 



(PPt) 

(°C) 

(ppm) 

(m) 

STATION 1 






15 May 

1.5 

7.6/7.4 

14.0 /14.1 



2 3 May 

2 June 

14 June 

27 June 

12 July 

4 August 
23 August 
18 Sept. 


5.5 

18.0 /18.0 

9.7 / 9.5 

1.0 

STATION 2 






15 May 

3.3 

7.1/7.1 

14.3 /14.5 



23 May 


5.5 

20.0 /19.0 

9.9 / 9.2 

0.8 

2 June 


5 




14 June 

27 June 


6.3 

25.3 


1.3 

12 July 

4 August 
23 August 
18 Sept. 


7.5/7.5 

< 

25.0 / 22.5 

6.9 / 6.5 


STATION 3 






15 May 

1.5 

6.9/7.1 

14.1 /13.8 



2 3 May 


5.8 

24.0 / 24.0 

9.3 / 9.8 

0.5 

2 June 


5 



0.6 

14 June 


5 

21.3 

7.8 

27 June 


6.5 

26.5 


1.1 

12 July 


7.0 

24.7 



4 August 


8.0 

25.8 

5.5 


23 August 


10.0 

27.2 

6.8 


18 Sept. 


13.0 





(continued) 



14 











TABLE 1. (continued) 



Depth 



Surface/Bottom 



Secci 


(m) 

Salinity 

Temperature 

Dissolved 0 0 

Depth 



(PPt) 

(° 

’C) 

(ppm) 


(m) 

STATION 4 









15 May 

3.6 

6.6 

/6.8 

15.1 

/14.6 




2 3 May 


5.2 


19.5 

/19.0 

11.4 / 

8.4 

0.5 

2 June 


6 







14 June 


5 


20.8 


7.5 


0.6 

27 June 


6.2 


28.0 




0.8 

12 July 


6.2 

/ 6.2 

24.7 

/23.5 




4 August 
18 Sept. 


7.0 


26.1 

/25.8 

7.4 / 

5.6 


STATION 5 









15 May 

3.0 

6.0 

/6.0 

15.3 

/15.3 




23 May 


5.8 


21.0 

/19.5 

9.5 / 

9.6 


2 June 

14 June 

27 June 

12 July 

4 August 
18 Sept. 


5.0 


21.0 


8.4 


0.9 

STATION 6 









15 May 

3.6 

5.8 

/6.5 

15.3 

/14.8 




2 3 May 


5.9 


19.0 

/19.0 

9.3 / 

7.8 

0.6 

2 June 

14 June 


5.5 


21.0 


8.0 


0.7 

27 June 


6.0 


28.2 




0.6 

12 July 


7.2 

/6.8 

25.2 

/24.4 




4 August 


6.2 


26.2 

/25.9 

5.9 / 

4.7 


23 August 


8.0 


28.8 

/26.6 

7.0 / 

5.4 


18 Sept. 


9.0 







STATION 7 









15 May 

2.4 

5.4 

/ 5.4 

15.3 

/15.3 




2 3 May 


5.5 


19.0 

/18.6 

8.2 / 

7.7 


2 June 

14 June 


5 


22.1 


7.2 


0.7 

27 June 


6.0 


28.3 




0.7 

12 July 


6.0 

/5.9 

25.2 

/25.8 




4 August 


6.0 


26.2 


5.9 



23 August 


8.0 


27.4 

/27.0 

6.8 / 

5.8 


18 Sept. 


8.0 








(continued) 


15 











TABLE 1. 


(continued) 


Depth 

(m) 


Salinity 

(ppt) 


Surface/Bottom _ 

Temperature Dissolved 0~ 
(°C) (ppm) 


Secci 

Depth 

(m) 


STATION 8 


15 May 

3.4 

5.0 / 5.0 

15.2 / 15.2 


2 3 May 


4.8 

18.0 / 19.0 

9.2/ 8.3 

2 June 


6 



14 June 


5 

22.0 

6.9 

27 June 


5.5 

28.0 


12 July 


5.8 

26.2 


4 August 


6.0 

26.5 

5.4 

23 August 


8.0 

27.8 / 27.0 

6.5 / 5.4 

18 Sept. 


7.0 



STATION 9 





15 May 

4.3 

4.4 / 4.2 

15.3 / 15.1 


2 3 May 


4.5 

18.5/18.5 

8.2 / 8.0 

2 June 


6 



14 June 


4 

23.0 

6.8 

27 June 





12 July 


5.0 

26.0 


4 August 


5.8 

26.5 

5.8 

23 August 


7.0 

27.8 27.0 

5.8 / 5.2 

18 Sept. 


9.0 



STATION 10 





15 May 

14.3 

3.3 / 3.3 

15.7/15.7 


23•May 


3.0 

19.0/19.0 

8.0 / 7.5 

2 June 


2 



14 June 


2 

22.5 

7.5 

27 June 





12 July 


2.2 

26.2 


4 August 


3.0 

26.9 

4.8 

23 August 


5.0 

27.5 

6.0 / 5.6 


18 Sept. 


0.6 

0.5 


0.4 


0.2 


16 













PROCEDURES 


Oysters used in the field experiment had to be obtained from 
a source that was free of phthalic esters, therefore, the natural 
oysters in the Chester River could not be used. Cultchless 
oysters with a height of approximately 3 cm were bought from 
Frank Wilde's Hatchery in Shadyside, Maryland. The oysters were 
distributed randomly over 15 trays with 96 oysters per tray. 

Trays were of stainless steel and measured 40 cm x 92 cm x 10 cm 
with a mesh of 2 cm x 2 cm. The lids were hinged by two rings 
through both lid and tray on one of the long sides and a stain¬ 
less steel clip hasp lock on the opposite side. Each tray was 
bridled by a 6.4 mm braided nylon line from each top corner that 
were joined about 1 m above the tray with a loop. Another 6.4 mm 
nylon line was fastened to the loop on one end and to the station 
on the other. Stations 8, 9 and 10 were suspended, but touching 
the bottom, from permanent structures; all other trays were set 
on the bottom with the line attached to either a buoy anchor 
chain, a ring on a day beacon, or to private docks. All trays 
had two bricks tied on edge to their bottom to hold the trays 
above the muck. 

Sampling Visits 


A total of nine visits were made to the study area. Eight 
visits were collecting and observation field trips after the sta¬ 
tions had been established on 15 May. These visits were made on 
23 May, 2 June, 14 June, 27 June, 12 July, 4 August, 23 August 
and 18 September. Not all stations were visited on each field 
trip due to mechanical breakdown of the outboard motor. Dates 
of visits and stations sampled are given below. 

Visit 1, 15 May--Was done from R.V. AQUARIUS and established the 
stations under the guidance of Dr. George Krantz. All sub¬ 
sequent visits were made by outboard runabout. 

Visit 2, 23 May--Twelve oysters were collected from every sta¬ 
tion except station 1 where the tray had been turned upside 
down with the lid open. Qualitative notes were kept on 
presence or absence of organisms associated with oyster com¬ 
munities. For hydrographic data see Table 1. 

Visit 3, 2 June--Visited stations 2, 3, 4, 8, 9 and 10 success¬ 
fully. Station 1 had been abandoned. Station 6 could not 
be found. Stations 5 and 7 had gotten so badly tangled up 
in the rusty metal of the day markers that the trays could 
not be brought to the surface for inspection and sample 
collection. 

Visit 4, 14 June--Successfully collected samples from stations 
3 - 10. 


17 



Visit 5, 27 June—Successfully collected oyster samples from 

stations 2, 3 , 4, 6, 1 and 8. Station 5 had been removed by 
vandals. Outboard motor trouble prevented sampling of sta¬ 
tions 9 and 10. 

Visit 6, 12 July--Collections were made from stations 2 , 3 , 4 , 6, 
7, 8, 9 and 10. 

Visit 7, 4 August—Collected from stations 3, 4 , 6, 1 , 8, 9 and 
10. Station 2 could not be retrieved by either grapnel or 
by diving. Efforts to locate this station's trays on 
subsequent visits failed. 

Visit 8, 23 August—Failed to bring up station 4 with either 

grapnel or diving. Samples were taken at stations 3, 6, 7, 

9 and 10. 

Visit 9, 18 September--Ended the observation period. One tray 
from each remaining two-tray station was taken out. The 
remaining trays were left to be collected at a later date 
so that oysters could be checked for phthalic esters after 
a full growing season. However, those trays tied to the 
clam buoys were taken up by D.N.R. and no identification 
of trays could be made. 

Controls—A control station was established on 15 May at the 
pier at Solomons. Sampling of the controls coincided with 
every visit to the Chester River stations. 

Oyster Scoring Procedures 


Oysters were brought into the lab for gross examination and 
preservation. Oyster samples collected from the tray stations 
were packed in wet paper towels and kept under refrigeration 
until they could be examined. Oysters were shucked by severing 
the aductor muscle at the left valve and displaying the oyster 
in its right valve with the left valve removed. Features de¬ 
scribed were shell height, new growth, color of the meats, 
condition ,(which will be further explained) and possible imper¬ 
fections. Most common imperfections were Polidora websteri 
infections, shell ulcers, muscle ulcers, and mud blisters. 
Tissue ulcers and bill obstructions were rare and no pea crabs 
Pinnotheres ostreum or sponge penetration by Cliona celata were 
found (salinity too low). 

Length and new growth measurements were recorded in mm, but 
imperfections and infestations were scored on a scale of 0 (not 
present) to 5 (omnipresent). The scoring of observed oyster 
condition (not to be confused with condition index (Cl) which 
is a totally different procedure) was also according to a scale 
of 0 to 5 but deserves elucidation of its criteria: 


18 









0 = Dead. 

1- = An oyster that is completely transparent is definitely un¬ 
healthy, all organs clearly visible (no gonad or glycogen). 
It does not fill its shell cavity but lies like a limp 
piece of clear gelatin on its valve. This oyster is mori¬ 
bund . 

1 = Same as above with slight nuances for improved scoring. 

This oyster may live. 

2 = Oyster not transparent but liver (digestive gland) clearly 

visible. Gonad or glycogen low and patchy. Does not fill 
its shell cavity well. Not moribund. 

3 = Average looking. Reasonable amount of gonad and/or glyco¬ 

gen. Liver barely visible. Mantle appears to be an active, 
well-functioning organ. If examined immediately after col¬ 
lection a crystalline style may be found. 

4 = Good looking. Enough "fat" to cover liver completely. 

Plumper than 3, therefore, fills its shell cavity shell. 
Mantle uniform rich color. Mantle edge active to stimu- 
lae. Healthy heartbeat. Solid crystalline style upon 
examination immediately after collection. 

5 = Excellent looking. The oyster appears cramped by its 

shell cavity. It bulges over the half shell with thick- 
layers of gonad and/or glycogen. Clean uniform coloration 
throughout. No deficiencies of any kind noted. 

Upon completion of this gross examination, the oyster meats 
were individually wrapped in cheesecloth together with a num¬ 
bered tag for later identification and preserved in Davidson's 
fixative. 

Data for new growth and mean condition score were compared 
using Student's t-test. 

Possible Point Source Studies 


A 96-hour toxicity study was performed at stations in the 
vicinity of possible point sources of pollution. These sources 
were the Chestertown Sewage Treatment Plant, the Tenneco chemi¬ 
cal plant which manufactures plasticizers and the Campbell's 
Soup Company where chicken carcasses are prepared and diced. 
Tenneco and Campbell's discharge into Morgan Creek while the 
Chestertown Sewage Plant discharges into Radcliffe Creek shortly 
before it joins the Chester River at the southern edge of Ches¬ 
tertown . 


19 



Stations— 


There were a total of six stations. The control station was 
located at least 1-1/2 km upstream from the Tenneco plant. This 
station was in a small tributary to Morgan Creek but the same one 
which carries the overflow from the discharge pond of the Tenneco 
Pond to Morgan Creek. 

Tenneco Pond located in the Tenneco discharge pond about 6 m 
away from the standpipe which is the point farthest away 
from the plant's point of discharge. Depth at this station 
was about 0.6 m. 


Tenneco Downstream—in the receiving stream from the Tenneco 
Pond about 350 m downstream from the pond. Depth at this 
station was 0.5 m maximum. No tidal influence was noted. 

Campbell Upstream—in Morgan Creek near the middle discharge 
pipe of the Campbell leaching fields in 0.8 m depth at low 
tide. Due to tidal amplitude, this station could only be 
checked once a day as at high tide the depth became 1.6 m 
or more. 

Campbell Downstream—also in Morgan Creek about 500 m from 

station upstream and located just upstream from Campbell's 
most downstream discharge. Tidal difference here was also 
more than 0.6 m, which often limited mortality checks to 
once a day. 

Sewage Plant—was located in Radcliffe Creek about 6 m down¬ 
stream from the sewage outfall. Tidal difference varied 
depth from 0.5 m to well over 1 m, which allowed only 
one inspection per day. 

Procedures-- 

Experimental species for this investigation were the golden 
shiner, Notemigonas crysoleucas and the crayfish, Procambarus 
acutis acutis . These species were chosen because they are 
native to the area, are easily available, and easily acclimated 
to new conditions and represent animals which are neither overly 
delicate nor particularly pollution resistant forms. Golden 
shiners were obtained from Green Valley Minnow Farms, Brogue, 
Pennsylvania. Crayfish were obtained by seining various creeks, 
ditches, and rivers on the Eastern Shore. All animals were held 
for a minimum of 2 weeks in sand-filtered Solomons well water. 
Temperatures were adjusted to those that were expected to be 
encountered in the field. Both species were fed Purina Trout 
Chow daily. Transportation to and from the laboratory was in a 

1400 1 tank aerated using tanks of compressed air. 


Test animals were contained in cages of 30.5 cm x 30.5 cm x 
91.4 cm with a 13 mm rebar frame covered with 3.2 mm mesh nylon 
netting. 


20 








The experiment lasted 96 hours. One cage with 30 golden 
shiners and one cage with 15 crayfish were put overboard at each 
station. Where possible, the cages were checked twice daily and 
dead animals were removed and preserved for future autopsy. 

Once a day the animals were fed Purina Trout Chow®. 

All survivors of the 96-hour experimental period were pre¬ 
served in Davidson's preservative for later autopsy. 

Hydrographic data, recorded at every visit, were temperature, 
salinity, dissolved oxygen and residual chlorine (see Table 2 for 
these data). 

Autopsy of Specimens-- 

The autopsy of golden shiners was divided into two parts-- 
external and internal features. External features included 
standard length, examination for damage such as broken, absent 
of excessive mucus covering; missing or deformed scales, abra¬ 
sions on body and/or fins; and afflictions such as fungus, dis¬ 
colorations, cysts and parasites. Internal autopsy examined 
general appearance, color, damage and foreign material on or in 
the buccal cavity, gills and gill arches; texture, size, color, 
content and abnormalities of stomach, internal and external 
intestinal lining, liver, gasbladder, gonads, spleen and adipose 
tissue. Any observations that did not fit into the prepared 
autopsy sheet were recorded under item "Other." All specimens 
were individually wrapped in a bag of cheesecloth containing an 
identifying tag and were placed in fresh Davidson's solution. 

Cages containing crayfish were checked at the same time as 
the cages with golden shiners. At that time dead crayfish were 
removed and preserved in Davidson's solution. All surviving 
crayfish were preserved at the end of the 96-hour experimental 
period. 

Autopsy of crayfish examined both external and internal 
features. External features such as injury, regeneration to ap¬ 
pendages and body, hardness of cephalothorax, color, affliction 
with fungus, discoloration, and bacterial infection were recorded. 

Internal features such as condition and foreign material in 
the gills and gill chamber, texture and content of cardiac and 
pyloric stomach, condition and appearance of heart, hepatopan- 
creas and gonads were recorded. 

After autopsy the specimens were wrapped individually in a 
cheesecloth bag that contained an identification tag and placed 
in fresh Davidson's preservative. 

Autopsy data were examined for correlations using contingency 
analysis with chi-square tests. 


21 


TABLE 2. HYDROGRAPHIC OBSERVATIONS FOR THE POSSIBLE POINT SOURCE STUDIES 



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22 


(continued) 












TABLE 2. (continued) 


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23 


1430 17.5 4 8.7 0.12 



















RESULTS 


Oyster Experiments 

Mortality over the period was low (see Table 3). The only 
station showing mortality noticeably higher than that of the 
control station is Station 10. This is the most upriver station 
and subjectively seemed initially to have lower concentrations 
of fouling organisms and more silt. The single event of six 
dying, between 2 June and 14 June, may be related to heavy runoff 
and smothering by silt. 


TABLE 3. MORTALITY OF OYSTERS DURING THIS STUDY 







Station 







2 

3 

4 

5 

6 

7 

8 

9 

10 

Control 

23 May 

0 

0 

0 

* 

0 

* 

0 

0 

0 

2 

2 June 

1 

0 

0 

* 

* 

* 

1 

0 

1 

1 

14 June 

* 

0 

0 

0 

0 

0 

1 

0 

6 

0 

27 June 

0 

1 

0 

It 

1 

0 

0 

* 

★ 

0 

12 July 

* 

0 

0 


0 

1 

°$ 

0 

0 

0 

5 August 

* 

0 

1 


1 

1 

1 

1 

1 

0 

Total 

1 

1 

1 


2 

2 

3 

1 

8 

3 


★ 

Trays 

were 

not 

examined 

• 






+ Lines 

cut 

and 

tray 

missing. 







| Tampered with but appeared intact 

In late spring and early summer, growth was mostly not 
significantly different from that shown at the control station, 
and there was no pattern of either increased or decreased growth 
(see Table 4). In July, all stations showed significantly slower 
growth than the controls while in August four out of seven sta¬ 
tions showed significantly slower growth. 


24 










TABLE 4. COMPARISON OF MEAN NEW GROWTH IN OYSTERS PLANTED IN CHESTER RIVER* 


O 

p 

-p 

G 

o 


cn 


G 

O 

•H 

■p 

0 

-p 

co 


00 


vo 


in 




ro 


r\i 


o 


ro 

CO 

o 

rx 

O' 

rH 


00 

ro 

o 

rH 

O’ 

• 


• 

• 

• 

• 

• 

VO 


CN 

N* 

VO 

00 

00 






x _ v 







rH 

rH 

CN 

• 

ro • 

CN • 


in o 

CN O 

00 

in 

r- co 

N* CO 


CN o 

H 1 O 






• • 

• • 

in 

2 

ro 2 

O' 2 


co o 

rH O 


-- 

'-- 

— 


— 












s 


rH 


00 

• 

CN • 

in • 


o o 

CN • 

o 

in 

cp co 

rH co 


in o 

O' CO 






• • 

• • 

in 

z 

ro 2 

ro 2 


in o 

rx 2 


v ' 

v * 



' * 




^^ 





Cn 

rH 

• 

00 • 

o • 

in 

ro cn 

CN 

o 

iH co 

in co 

in co 

vo o 

00 o 


o 


H 1 2 

m 2 

in o 

vo o 


—" 

-- 

"— 


--- 

"—" 





,_^ 





♦ 

CN rH 

CN • 

H 1 iH 

r- rH 




Cn O 

cp cn 

O O 

rH O 




• • 







in o 

vo 2 

VO o 

vo o 




-' 

'—" 

■—' 

'— 









rH 


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rH 


O 

o 


oc • 

r- • 

CO o 

CN • 

rH 

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o co 

vo co 

ro o 

cn CO 

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• 






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o 


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O’ o 

rx 2 


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• 

r- cn 

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r- 

in 

vo o 

in co 

O CO 



in 

2 

O 

ro 2 

vo 2 






v ' 








^^ 



00 

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o • 

ro rH 

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00 

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cn co 

co co 

O CO 

00 o 

O CO 

in 

s 

2 

N< 2 

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in o 

CT1 2 



' 1 

' " 

v —’ 

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' 












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rH 

rH 

in 

• 

in • 


• 

o o 

o o 

CN 

in 

r- in 


rH CO 

in o 

CN O 

• 

• 

• • 





VO 


CO 2 


ro 2 

O' o 

H 1 O 


N ** 












■P 







0 



0 

0 

0 

>i 

3 



G 

c 

G 

rH 

CP 

0 


3 

3 

3 

3 

3 

2 


P) 

►0 

•0 

*0 

< 

ro 


CN 

H* 

r- 

CN 

in 

CN 



rH 

CN 

rH 



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-P 

0-H 

0-H 

•P rtf 
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£ 0 

fd ^ 

g 

If) 

w 0) 

0 4J 

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L U 

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c 

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rd 

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o w 

p . 
(3^2 

Q) 

P . 
0 cn 
G 

w td 
CU 0 

W g 

0 

4^ rH 


-P 

c 

<u 

p 

fd 


o 

p 

-p 

g 

o 


CP o 

5 e 

•H O 
P 

W M-i 
P 

0 >, 

*9 1-1 

§ ^ 

2 td 

o 

•H 
• M-4 

E *H 

6 G • 
Cp in 
•p o 

0 • 

o 

13 p 
<D <u 

W 14-1 
w i4_i 
0 -H 

P T5 

a 

x +J 

CD o 

G 
CD 

£ 0 
<T3 eg 

in 
0 
3 


G 

■H 


G 

(d 

.G 

-P 

P 

CD 

Cp 

P 

fd 

r—I 


CO 0 

G G 
fd fd 
0 0 
g E 


0 

> 


25 
















There were few significant differences from the controls in 
observed condition scores (see Table 5), however there is a pat¬ 
tern that lower salinity stations accounted for most of these 
differences. In all cases but two, where significant differences 
occurred, the average condition scores were higher than that of 
the controls. These aberrant stations are Stations 9 and 10 
which are the two most upriver stations. 

Condition scores are affected by the seasonal condition 
changes that normally occur in oysters and only partially reflect 
(except in extremes) the oyster's health. Condition scores are 
based on the amount of solid color and how well the oyster fills 
its shell. The buildup of gonad in the spring and the accumu¬ 
lation of glycogen in the fall tend to elevate the score. Con¬ 
versely, low scores should be expected at the end of summer when 
oysters are completely spawned out and have suffered through 
unfavorable high summer temperatures that adversely affected 
their pumping and feeding. Also, oysters that have come through 
a hard long winter and spring with low temperatures and low 
available food have used up a great deal of their reserves (gly¬ 
cogen) and therefore will score low. Still the health of these 
oysters would not be as different as the two separate scores 
seem to indicate. Salinity can also influence oyster condition. 
Oysters in low salinity upriver areas that often warm up early 
will start their gonad development earlier than oysters in the 
more saline downriver or bay areas that warm up slower. Freshets 
often cause upriver salinities that are too low for spawning at 
spawning temperature. These oysters will not spawn and in the 
fall convert their gonad material immediately into glycogen with¬ 
out going through a "summer slump." These oysters may score high 
all year round. However, during the same period the oysters 
downriver are likely to spawn because the salinity is favorable 
at spawning temperatures and enter the fall in low condition. 
After temperatures drop (usually later than upstream), they will 
feed effectively again and increase their score until the water 
temperature is 5°C when they cease feeding and rely on their re¬ 
serve food which decreases their score. Thus, oyster scores 
should be evaluated with season and location in mind using as 
many individuals as possible. It is a subjective comparative 
judgment and should be performed as much as possible by one 
person to eliminate differences between workers. 

Associated organisms and time of occurrence are given in 
Table 6. When trays were inspected on 12 July, there were no 
live commensal organisms present at Station 9, and at Station 
10 the shells were almost clean except for a few live barnacles, 
bryozoan colonies and amphipods. During inspection of oysters 
on 5 August, it was noted that very few commensal organisms were 
present at Stations 6, 7, 8, 9 and 10 and that the shells were 
cleaner than before. On 23 August, it was noted that algae and 
barnacles were dying and decomposing at Stations 6 and 7, and 


26 


TABLE 5. COMPARISON OF MEAN OBSERVED CONDITION SCORES IN OYSTERS PLANTED 

IN CHESTER RIVER* __ 


G 

o 

•H 

-p 

fT3 

■p 

CO 


rH 

rH 


00 

CD 

CD 

o< 

ct\ 

0 

ro 


CM 

O’ 

in 

r- 

O' 

p 

• 


• 

• 

• 

• 

• 

-p 

ro 


ro 

ro 

ro 

ro 

ro 

c 








o 








u 















rH 


CD 

• 

00 • 

cn rH 


in • 

00 o 

o 

O’ 

01 

rH in 

on o 


cp co 

CD O 

rH 






• • 

• • 


ro 

2 

ro 2 

CM o 


ro 2 

o' o 



-- 

-- 

+ 


-- 

- 












.—x 



,—■* 

rH 


in 

• 

ro • 

in • 


rH in 

ro o 


CM 

CO 

CM CO 

00 CO 


CM O 

oo o 

cp 






• • 

• • 


ro 


ro 2 

<o 2 


ro o 

O' O 



'— 

— 



+• —- 

- - 













rH 



rH 


rH 

• 

00 • 

ro o 

r- 

00 iH 

uo o 


rH 

CO 

ro CO 

ro o 

CM o 

C' o 

r- o 

00 









ro 

2 

ro 2 

•O' o 

O' O 

O' o 

O O 



'—’ 

--- 

1 — 1 

'—’ 

1 ■ 

' ' 













rH 



•H 





O' O 

ro • 

ro • 

■O' o 





ro o 

oo co 

CD CO 

r- o 





o' o 

ro 2 

ro 2 

O’ o 






s * 


'' **’ 






^^ 





• 


m • 

O’ • 

ro • 

r- • 


o 

CO 


CO CO 

r- co 

r- co 

CD CO 

CD 

• 

• 







ro 



ro 2 

ro 2 

co 2 

ro 2 





' s “^ 


"" *” 






# __ 








00 • 








CD CO 








• • 








ro 2 








'' ^ 










4 __ 

4 __ 


rH 

• 

oo in 

<T« • 

rH • 

o • 

CM • 


o 

CO 

CD o 

r- co 

CO CO 

CO CO 

o' co 

o' 









ro 

2 

ro o 

ro 2 

ro 2 

ro 2 

ro 2 



" * 


V * 

N ^ 

' ** 





^^ 






ro 

• 

o • 


CM • 

00 • 

CD • 


ro 

CO 

CM CO 


r- co 

uo co 

O CO 

ro 

• 

• 

• • 






ro 

2 

ro 2 


ro 2 

ro 2 

o’ 2 



" 

' ' 


' ’ 





„_ N 







ro 

• 

00 • 


m • 




ro 

co 

rH CO 


in co 



CM 

• 

• 

• • 


• • 




ro 

2 

ro 2 


ro 2 






V ’ 












-P 








CO 




CD 

cu 

0 

>i 

p 



G 

G 

G 

rH 

Cp 


rd 


P 

P 

P 

P 

P 


S 



P) 

•"3 

•o 

< 


ro 


CM 

O' 

r- 

CM 

m 


CM 



rH 

CM 

rH 



I 

-p 


G 


rd 


0 


•H 


mh 


•H 


G 


CP 

• 

•H l 

uo 

co 

o 


• 

p 

o 

<D 


ip 

G 

ip 

rd 

•H rG 

Ti 

■P 

-p 

P 

0 

0 

G 

CP 


P 

0 

0 

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CO 

in 

G 

0 

rd 

P 

0 

rH 

£ 

0 


> 

<u 


CO 

>1 

0 

-P 

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•H 

-p 

rH 


•H 

-p 

a 

rd 

rd 

.G 

A 

-P 

0 


P 

co 

CU 

<1) 


•H 

0 

-P 

0 

•H 

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rH 


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u 

Xi 

•H 

rd 

TS 

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C 

0 

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P 


CU 

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0 

• 

P 


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CO 

• 

0 

CO 

co 

G 

0 

0 

.G 

0 

-P 

e 

G 


0 

rH 

P 

0 

0 

p 

CU 

-p 


G 

G 

0 

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O 

CO 

e 

P 

0 

0 

p 

rQ 

mh 

P 


2 

rH 


* 


• 

CD 

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rd 

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o 

p 

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g 

o 

u 

c 

rd 

£ 

-p 

p 

0 

2 

o 


>1 

r—I 

-P 

c 

rd 

u 

•H 

4H 

•H 

a 

Cp 

•H 

CO 


CO 

0 

3 

i—I 

rd 

> 


4 - 


27 



















TABLE 6. ASSOCIATED ORGANISMS BY STATION 




00 



ID 

00 

VO 

vo 

£ 









0 


VO 


00 

■^r 

VO 

ID 


•H 









-p 

•^r 

CO 

VO 

VO 

CM 

co 


CO 


vo 


ID 


CO 


CM 


CO CO 


00 

vo 

UO 


CO 


rr 

co 

•. V 

(N .H 


VO 


C" 




00 


00 








vo 



vo 

VO 







V 



ID 


vo 

LD 

uo 

uo 

vo 







•*. 

■rtf 


ID 

■^T 



CO 


CO 


CO 


CO 


r- 

VJO 


CM 


CO 


00 




00 

co 


vo 



■ V. 






vo 



uo 







vo 

CM 

uo 

vo 

vo 

■^r 



** 




ID 

rH 


UO 

UO 

CO 




vo 















•H 













,— 





•H 













CO 





m 






0 







•H 





•H 






fd 







> 





P 













p 





P 


in 




pH 







3 





3 


3 




fd 







U 




• 

X 


XS 









CO 


0) 

m m 


fa 



•H 




£ 





0) 


P 

TS t 


CO 

0 


fa 


•H 


0 





-p 



c 

0 



3 


3 


O 


0 





<D 


m 

fa fa 


CO 

0 


0 


0 


p 





3 


•H 

•r— 

•H 


0 

fa 




0 


CO 





X3 


-P 

X 

x 


-p 

0 


0 


X} 







o 


£ 

fa fa 

fa 0 

XI £ 


0 




in 

<D 

0 ) 



Po 


0 

£ 

£ 

£ 

£ 

3 3 


-P 


3 


3 

O 

3 



1—1 


TJ 

3 3 

•H 

O 

P fa 


u 


>i £ 

0 

0 

3 

CD 

m 


o 


0 



p 

£ 

O 0 

X 

0 

0 

X 0 

£ 

-P 

-P 

r—i 

£ 

m 

fa 


Po 

TS Td 

X 

0 

P 

3 

£ 

0 

0 0 

m 

£ 

-P 

3 

3 

TS 


CO 

XI 

•f 

•H 

CO 

0 

^ X 

P 

•H 

rH 

Cn 0 

•H 

0) 

0 


0 

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t 

I—1 

U 

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3 

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O rH 

U 

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£ 

£ 

i—1 

£ 

N 

0 

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0 

3 

ft 

fa 

co 

1—1 

X -H 


rH 

3 

T5 XI 

3 

fd 


£ 

0 

P 

0 

CO 

P 


0 

CO 

fd 

-P X 

0 

fd 

£ 

0 0 

to 

rH 

3 

o 

>i 

T5 

P 

CO 

CQ 


P 

3 


£ fa 

3 O 

P 

X O 

p 

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p 

P 

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3 

3 

—' 

ft 

0 

p 

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rH 

-- 

3 

3 

o 

fa 

in 

PQ 

PQ 

PC 

2 

S 


u 

u 

O 


X 

PQ 


PQ 

2 


28 


(continued) 


















TABLE 6. (continued) 


3 

o 

H 

4-> 

0 

X 

CO 




00 


00 



00 

00 








VO 



m 

r~ 

00 







in 


00 

ro 

VO 

VO 





*h 


ro 


r- 

CM 

CM 

in 





00 























00 

VO 

00 










r- 

ro 










00 

ro 

(N 

VO 







ro 


r- 

Csl VO 

rH 

<n 


VO 

IT) 



00 





00 









vo 

00 



ro 

r- 




•k 


h. 

*► 


in 

r* 


in 

CN 

VO 






%k 


ro 


vo 

ro vo 

ro 

1 —1 

in 


00 















00 



r- 






00 




00 

r- 




vo 



vo 





r- 

in 


in 



in 




in 



in 




















r" 



ro 

ro 

vr 

ro 


vo 


vo 



ro 



ro 

vo 
































•H 













,—> 





•H 













w 





in 






0 







•H 





•H 






0 







> 





P 






tr> 







P 





P 


in 




i—i 







3 





0 


3 




id 







o 




• 

X 


T5 









in 


0 

m 

w 


CL 



•H 


,—V 


3 





0 


p 

Ti 

TS 


in 

in 


CL 


•H 


0 





-P 



0 

0 



3 


0 


U 


0) 





0 


in 

CL, 

CL 


in 

0 


in 


0 


p 





0 


■H 

•H 

•H 


0 

CL 




0 


CP 





£ 


-P 

X 

X 


-P 

0 


in 


X 







o 


G 

CL 

CL 

CL 0 

X 3 


0 




in 

CD 

0 



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0 

£ 

£ 

£ 

G 

0 0 


X 


0 


3 

O 

id 



rH 


t 

id 

0 

•H 

O 

P CL 


u 


>i £ 

CO 

0 

3 

CP 

in 


0 


0 



P 

£ 

O 0 

X 

0 

in 

X 0 

£ 

X 

X 

rH 

3 

in 

a 


>i 

TJ 

t 

X 

0 

P 

0 

3 

0 

0 0 

0 

3 

X 

id 

id 

t) 


0 X 

•H 

•H 

m 

0 

TS X 

P 

•H 

X 

CP 0 

•H 

0 

CD 


0 

•H 

T3 

rH 

O 

P 

x 


0 

•H X 

u x 

o 

•H 

3 

£ 

rH 

3 

N 

0 

•H 

0 

0 

0 

CL 

m 

rH 

X -H 


rH 

0 

TJ X 

0 

<d 


£ 

o 

p 

0 

in 

P 

G 

0 

m 

03 

X X 

0 

fti 

3 

0 0 

Cr 

rH 

id 

0 

>i 

TS 

p 

in 

CQ 

G 

P 

0 

CH 

3 PH 

3 

U 

P 

X u 

p 

•H 

CD 

u 

p 

>1 

0 

3 

— 

0 

0 

p 

■— 1 

0 

rH 

-- - 

0 

0 

o 


CO 

m 

m 

K 

2 

£ 


O 

U 

O 


X 

DO 


DO 

2 


29 


(continued) 



















_ TABLE 6. (continued) __ 

_ Comments ___ 

Station 2. Very few associated organisms throughout study; lost, not sampled 

after 27 June. 












03 


p 






TS 






0 


d 





CO 

G 


CO 0 




P 







-P 

d 


P d 



1 

O 


0 





G 



G Cn 



G 

0 


co 


»— H 



0 

tn 


0 rH 



•H 

a 


G 


rH 



P 

G 


p d 




CO 


0 


d 



P 

•H 


p 



P 

G 


03 





P 

>i 


P r-- 



d 

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c 



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T5 


u 






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0 






G 



CO 

p 


P 


TS 



1—1 

0 


rH 0 



E 

0 


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d 

d 


d tH 



CO 

G 


G 


CD 



TS 

Cn 


03 P 



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P 


-P 



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rH 


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G 

• *. 


P 


o 



P 

d 


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d 

00 


g 


0 






0 



Cn 



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0 

Tl 


0 CO 



P 

G 


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CO 



rH 

G 


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O 

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XI 

d 


X ’H 




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CO 


p p 



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rH 


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d 

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CO 


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MH 

d 


Mh r* 



o 

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MH 





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G 


G 



MH 

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CO 



P 

p 


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03 




• %» 


G 




d 


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G 

£ P 


f- 


0 



MH 

X! 


mh d 


d 

0 

d 




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0 



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ro 



ID 


ID 


t>i 


ID 


O 



0 

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0 




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0 



CO a 



CO CO 


CO 

G 

•H 




a. 



P -H 

G 


P G 


G 

d 

CO 


in 


CO 



d P 

0 


d o 


O 

0 

G 



a 

G 



O P 

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O *r| 


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rH 

0 


co 

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0 

P 


0 P 


P 

0 

03 


G 

p 




X! G 

U 


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p 

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0 

0 


0 


0 

0 

G 


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P 



CN -H 

C, 


cn a 


a 

p 

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P 

G 




P 

CO 


CO 


CO 

0 



O 

O 

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30 










and fouling communities were still sparse at Stations 6, 7, 8, 9 
and 10. By 18 September fouling communities seemed back to nor¬ 
mal at all stations inspected; that is, all stations but 2 and 5. 

Golden Shiner Experiment 


Mortality was high at all stations including the controls. 
Figure 3 is a graphic representation of mortality over time. 
Excess mortality, that is mortality of controls minus mortality 
of experiments, is shown in Figure 4. Ranked in descending order 
of total survivorship the stations are Tenneco Downstream and 
Control, with Tenneco Pond, Campbell Upstream, and Campbell Down¬ 
stream all having the same survivorship and with The Sewage Plant 
having the lowest survivorship. Sewage Plant station is unique 
in the excess mortality curves in that all nonzero values are 
negative, that is mortality was higher than control mortality at 
all times. 



> 

QC. 

z> 

CO 



Figure 3. Mortality of golden shiners, by station, over the 
96-hour observation period. 


31 











SURVIVORS 



> 

CXL 

ID 

CO 




Figure 3. (Continued) 



32 













10 







Figure 4. Excess mortality of golden shiners, by station, during 
96 hours of observation. This is calculated by sub¬ 
tracting mortality of experimentals from that of the 
controls. Negative values indicate mortality more 
than that of the control. 


33 
















Autopsy data showed tho following to be significantly cor¬ 
related with death at one or more stations (see Table 7 for a 
presentation of these data); fungus on body or fins; watery fat 
in the mesentery and on the surface of the intestines; foreign 
material on the gill surface; and hard, white crystalline nodules 
attached to the body or fins. Comparing stations for frequency 
of occurrence of each of these characteristics using nonsurvivors 
only showed that no location had significantly higher or lower 
frequency of watery mesenteric fat; Campbell Upstream and Camp¬ 
bell Downstream had significantly higher frequency of occurrence 
of fungus; the Control Station showed a significantly higher fre¬ 
quency of foreign material (apparently silt) on the gills and 
Sewage plant, Tenneco Pond and Campbell Downstream showed sig¬ 
nificantly higher frequency of white nodules. See Table 8 for 
levels of significance of these characteristics. 


TABLE 7. 

RELATIONSHIP OF 

NONSURVIVAL 

TO AUTOPSY 

DATA* 


Station 

Fungus 

White 

nodules 

Foreign 
material 
on gills 

Watery 

mesenteric 

fat 


Control 

N.C. t 

N.C. t 

0.05 

0.05 

Campbell Upstream 

0.005 

N.C.t 

N.S. 

N.S. 

Campbell Downstream 

0.005 

N.S. 

N.S. 

N.S. 

Tenneco Pond 

N.S. 

N.S. 

0.05 

N.S. 

Tenneco Downstream 

0.05 

N.S. 

N.S. 

N.S. 

Sewage Plant 

N.S. 

0.01 

N.S. 

0.01 


Numbers are probability values calculated using contingency 
analysis. N.S. indicates that probability was greater than 
0.5. 

t 

Not calculable because of zero expected values. 



34 







TABLE 8. COMPARISONS BETWEEN STATIONS FOR CHARACTERS 

RELATED TO NONSURVIVAL* 




Campbell 

Campbell 

Tenneco 

Tenneco 

Control 

Upstream 

Downstream 

Pond 

Downstream 



Fungus 




Campbell Upstream 

0.005 





Campbell Downstream 

0.005 

N.S. 




Tenneco Pond 

N.S. 

0.005 

0.005 



Tenneco Downstream 

0.025 

0.025 

0.005 

N.S. 


Sewage Plant 

N.S. 

0.005 

0.005 

N.S . 

N.S. 



Foreign 

l material < 

on gills 


Campbell Upstream 

0.05 





Campbell Downstream 

0.005 

N.S. 




Tenneco Pond 

0.005 

N.S. 

N.S. 



Tenneco Downstream 

0.005 

N.S. 

N.S. 

N.S. 


Sewage Plant 

0.005 

0.05 

N.S. 

N.S. 

N.S. 



White nodules 



Campbell Upstream 

N.S. 





Campbell Downstream 

0.025 

0.025 




Tenneco Pond 

0.005 

0.005 

N.S. 



Tenneco Downstream 

N.S. 

N.S. 

N.S. 

N.S. 

N.S. 

Sewage Plant 

0.005 

0.005 

0.005 

0.025 

0.005 



Watery 

mesenteric 

fat 


Campbell Upstream 

N.S. 





Campbell Downstream 

N.S. 

N.S. 




Tenneco Pond 

N.S. 

N.S. 

N.S. 



Tenneco Downstream 

N.S. 

N.S. 

N.S. 

N.S. 


Sewage Plant 

N.S. 

N.S. 

N.S. 

N.S. 

N.S. 


* 

Numbers are probability values calculated using contingency 
analysis. N.S. indicates that probability was greater than 
0.05. 


35 












Crayfish Experiments 

No significant mortality was seen at any station, and 
autopsies failed to disclose any obvious difference that could be 
related to treatment between the crayfish that died during the 
experiments and those that survived. 

DISCUSSION 

Oyster Experiments 

Since one of the principal objectives of these studies was to 
identify possible causes of oyster mortality in the Chester River 
since 1974, it is unfortunate for the study that no large-scale 
mortality occurred at any of our stations. Observations by sci¬ 
entists, watermen and representatives of the Department of 
Natural Resources have noted that oysters that die in the "mor¬ 
tality areas" of the Chester River have unusually clean shells 
showing little or no fouling. We did note a dieoff of fouling 
organisms. This dieoff was first noted at the two most upriver 
stations on 12 July and coincided with a fishkill in the river. 
Our personnel noted that the fishkill was a widespread phenomenon 
with dead fish, mostly catfish and carp occurring from near the 
mouth of the river to well above the mouth of Morgan Creek. It 
might be speculated that the two phenomena are related in some 
manner. 

The dieoff of fouling organisms was never detected downstream 
below Station 6 (clamline buoy A off Corsica Neck) and seeming 
full recovery was noted by 18 September. An examination of Table 
3 will show an interesting pattern of deaths. If we ignore the 
unusual high mortality at Station 10 on 14 June, which may be re¬ 
lated to high runoff and silt, 6 of the 13 remaining mortalities 
occurred during the period of the fouling organism dieoff and at 
the stations where commensal organism dieoff was noted. It may 
be that the oyster dieoff did occur during the study, but for 
some reason this was an unusually mild year for it. 

An examination of Table 4 shows that summer growth rates of 
our oysters placed in the Chester River were significantly lower 
than that of the control oysters held in the Patuxent River. 

This reduced growth rate coincides with the period of time when 
the dieoff of fouling organisms occurred. 

Table 5 shows that oysters in the four most upriver stations 
had significantly higher observed conditions scores. These may 
well relate to salinities being too low for spawning. These 
oysters may simply have never lost gonadal material or glycogen. 

One possible hypothesis for the dieoff has been suggested by 
Donald Heinle (personal communication). He suggests that heavy 
loading of the system with organic material will cause the water 


36 


to go anoxic or nearly anoxic at depth, and under such conditions 
carbonic acid is produced which will dissolve the outer layer of 
the oyster shell; thus, producing the "scrubbed clean" look of 
the oysters. He has seen this occur in estuarine waters and in 
the laboratory despite the heavy buffering of the salts present. 
If in addition to producing carbonic acid, anoxic conditions hold 
for a long period when the oysters were otherwise stressed, then 
mortality would occur. This hypothesis is consistent with the 
data and cannot be disproved as our sampling periods were far 
enough apart that we might not have detected the event causing 
the anoxia. 

Golden Shiner Experiments 

Table 7 indicates that different stations show different cor¬ 
relations between nonsurvivorship and four factors noted during 
autopsies. Both Campbell stations and Tenneco Downstream had 
significantly higher infection percentages of fungus. The fungus 
appeared to be Saprolegnia sp. but was not identified. 

Saprolengia is not an obligate parasite. Usually it occurs as a 
saprophyte. Its abundance at these stations indicates high con¬ 
centrations of organic material. 

The white nodules are problematic. Their identification is 
unknown. They are white, translucent, hard, brittle and sub- 
spherical to ovoidal to rounded irregular in shape. Largest 
diameter is less than 1.5 mm. They occur in patches usually on 
the ventral half of the body or on the fins. They are firmly 
attached to the scales, fin rays or opercle but can be broken 
loose by scraping. None were noted in specimens which had sur¬ 
vived nor in any specimen from Campbell Upstream or the control 
station. It was thought that these might be the results of an 
interaction between the calcium of the boney scales and fin rays 
and some ingredient in Davidson's fixative. Arguments against 
this are numerous. The crayfish have as much or more calcium in 
their exoskeletons than the fish scales, but no crayfish was ever 
found with one. The placement on the body in discrete patches 
argues against any random process, and the fact that there were 
significantly more or less of these at certain stations adds 
doubt to any simple explanation. 

Foreign matter on the gills irritate the gills, cause stress 
symptoms and synergistically increase toxicity of other sub¬ 
stances.* It is therefore conceivable that the foreign material 
noted on the gills of fishes that died at the control station 
was a contributing factor to their death. In addition to silt 
particles on the gills, fibers resembling those from toilet paper 
were noted on one fish's gills. This indicates that untreated 
domestic sewage may find its way into the stream above our con¬ 
trol station. 

* Wilber, Charles G. 1976. The biological aspects of water pollu¬ 
tion. Charles C. Thomas, Publisher, Springfield, IL, 296 p. 


37 





What we have referred to as "watery fat" is a type of fatty 
tissue that has a flabby look and a semiliquid consistency. It 
appears under low magnification to be translucent and to have 
inclusions of water or other liquid free in it. This fat was 
found in both surviving and dead specimens but a significant cor¬ 
relation between it and the nonsurvivorship was noted at the con¬ 
trol station and at the sewage plant station. No mention of this 
kind of fat in the literature was found, but it was speculated 
that the presence of watery fat may be an early symptom of 
stress. In prolonged periods of high energy expenditure and 
during starvation, body fat is mobilized to act as an energy 
source. If lipids are used up, fat cells may lose their turgor 
producing this soft,gelatinous texture. 

The fact that dead fish from different stations have very 
different patterns of correlations with these factors indicates 
that there are probably multiple causes of mortality and that 
each station has its own unique combination of factors. In 
short, no single cause of mortality was found nor was it thought 
that the mortality noted in these experiments is necessarily re¬ 
lated to the cause of oyster mortality downstream. 

Crayfish Experiments 

Mortality was too low and autopsy data failed to find any 
changes which could be related to treatments. It seems that this 
species of crayfish is hardier than the golden shiners. The mor¬ 
talities occurred mostly with crayfish that had moulted and that 
were killed then by other crayfish. 


38 


SECTION 5 


PHTHALATE ESTERS AND RELATED CHEMICALS IN THE CHESTER RIVER 


INTRODUCTION 

The immediate purpose of this work is to assess the potential 
for bioaccumulation of alkyl phthalates as a way to explain the 
recent past mortality of oysters in the Chester River. Our 
starting approach involved dual plans to use both gas and liquid 
chromatography. Careful technique is needed to avoid contamina¬ 
tion by the ubiquitous presence of the phthalate esters in the 
environment, including their reported presence in the chemical 
laboratory. Our own facilities proved to be no exception. 

Two main problems needed to be solved: positive errors due 
to contamination during the measurement process, and negative 
errors due to incomplete extraction. Hopefully, both problems 
have been solved. Several models were considered for influx of 
alkyl phthalate waste from the Tenneco plant. These considera¬ 
tions were used to guide the sample selection and interpretation 
process. In brief, our overall approach emphasized measurement 
reliability, it narrowed the number of samples for analysis, and 
it gave a conceptual base to guide future work. Alkyl phthalate 
contamination in the Chester River clearly includes di(2- 
ethylhexyl)phthalate (DEHP). Our finding of massive contamina¬ 
tion in Tenneco Pond is consistent with the pollution conse¬ 
quences that are consistent with the existing discharge permit 
which has been approved by the State of Maryland. 

Geographic and Background History 

The Tenneco factory is sited on a tributary of Morgan Creek 
approximately 7 miles (10.6 km) upstream from where the creek 
empties into the Chester River. The location is shown in Figure 
5. The oyster mortality starts 12 miles (19.2 km) downstream 
from Morgan Creek toward the mouth of the Chester River. The 
mortality involved a downstream progression. Tne progressive 
movement is evident in Figure 6, which shows the status of 
mortalitities in oyster beds monitored in 1970-1975 (see 
Appendix A). The progression of the total mortality zone (0 
percent live oysters) is seen by comparing the 1973 and 1975 
results where the mortality front is seen to advance 5.2 km in 
the downstream direction. 


39 






Chester River Bridge at Chestertown 


Figure 5. Map of Chester River illustrating sample sites. 

Site coordinates are given in Table 18. 

* 


40 






T 



Figure 6. Downstream progression of oyster mortality. 


1 . 

Northwest 

17. 

Bailey 

33. 

Town Point 

2. 

Melton Point 

18. 

King's Creek 

34. 

Holton Point 

3. 

Booker Wharf 

19. 

Wilson's Point 

35. 

Oldfield 

4. 

Hollyday 

20. 

Eagle Point 

36. 

Robin's Cove 

5. 

Haddaway 

21. 

Island Point 

37. 

Chester River Mid 

6. 

Shippen Creek 

22. 

Davis Creek 


dleground 

7. 

Mummy's Cove 

23. 

Drum Point 

38. 

Bluff Point 

8. 

Deep Point 

24. 

Boathouse 

39. 

Hell's Delight 

9. 

Sheep 

25. 

Sand Thistle 

40. 

Bay Bush Point 

10. 

Commegy's Bight 

26. 

Hudson 

41. 

Piney Point 

11. 

Emmory Hollow 

27. 

Nichols 

42. 

Belts 

12. 

Spaniard Point 

28. 

Limekiln 

43. 

Durdin 

13. 

Cliff 

29. 

Willow Bottom 

44. 

Horse Race 

14. 

Ebb Point 

30. 

Possum Point 

45. 

Carpenter Island 

15. 

Ware 

31. 

Ship Point 

46. 

Black Buoy 

16. 

Phillip 

32. 

Emory Wharf 

47. 

Hail Creek 


- - 



49. 

Hail Point 





49. 

Poplar 

(From: Meritt, D. W. 

1977, Univ. Md. Ctr 

. Environ. Estuarine 

Studies Spec. Rpt. 7 

• ) 

41 












The discharge of any toxic chemical might account for the 
mortality pattern. The Tenneco plant waste discharge is one 

possibility based on circumstances that will be considered here. 


The Tenneco plant became operational in 1959. Esterification 
equipment was acquired in 1965. A process for easte treatment by 
bacterial digestion was installed in 1968. At this time, a dam 
was located across the water flowing from the Tenneco plant. As 
a result, a small pond (Tenneco Pond) was formed with 40 x 10^ m^ 
(10 acres or 4 hectares) of surface area. 


Hurricane Agnes occurred in May of 1972 bringing 16.0 cm of 
rainfall in one day (Palmer 1972) to the plant area which was 
flooded. The contents of a waste collection tank were washed 
out. It was estimated that 50-100 gallons (190-380 L) of organic 
waste liquid were washed out into Morgan Creek and then into the 
Chester River. It is possible that the washed-our chemicals 
could be causally related to the oyster mortality. (It is also 
possible that the effect might be the result of one or more other 
factors, including the continual discharge of unidentified organ¬ 
ic compounds which reside after waste treatment, agricultural 
runoff, a "natural phenomenon," etc.). Various pathways for the 
transport of phthalate esters from the Tenneco plant to the oys¬ 
ter mortality area are now discussed. 

The Upper Bay Survey (Palmer, Schubel, and Cronin 1975) 
makes it clear that sparingly soluble organic compounds are speci¬ 
fically associated with the finely has been established with ob¬ 
servations that the presence of such organic compounds is 
precisely correlated by the octanol-water partitioning model, and 
the adsorption of neutral organic compounds by sediment has been 
strongly correlated to the weight fraction of organic carbon 
(Karicknoff et al. 1979). This finding is consistent with the 
known tendency of humic acid to adsorb such compounds rather 
tenaciously. Schnitzer's work (1972) suggests possibly irrevers¬ 
ible adsorption of phthalate esters by clay soils, although there 
is some controversy on the significance of that finding. Regard¬ 
less, the known distribution of neutral organic compounds in the 
marine ecosystem includes biomagnification by life in the marine 
environment, and a corresponding adsorptive magnification effect 
into the natural clay-organic complexes present in marine sedi¬ 
ments. Since dioctylphthalate (DOP) was observed early in the 
study as a conspicuous and major component in Tenneco Pond sedi¬ 
ment, it is important to show whether or how that material would 
be transported from the Tenneco factory to the clay-rich sedi¬ 
ments in the Chester River. These latter sediments according to 
the Chester River Report (Palmer 1972) occupy the deeper terrace 
portions of the river floor and the transitional channels between 
nearshore and the main river channel. Taken in perspective, these 
are the most critical samples where investigation can be focused. 
Such sediments are widely recognized for their ability to adsorb 
compounds for long periods of time, geological epoch in many 


42 


instances. As a result, the clay-humic structure contains a 
historical reservoir of many compounds deposited through natural 
or anthropogenic influx. In the present study where no biologi¬ 
cal specimens were preserved, the key to the retrospective anal¬ 
ysis of the past oyster mortality would have to told by the 
sediment composition. 

DISTRIBUTION PATTERNS 

Three factors which may modify the distribution patterns of 
organic distribution in the Chester River as as follows. 

a. The downstream dilution effect . The width of the 
Chester River grows from roughly 0.5 km at Chestertown to 1.6 km 
near the midway region close to Spaniard Point to 5 km at the 
mouth. This suggests that the sediments, behaving as a pollu¬ 
tant sink, may become more dilute in the pollutant as one proceeds 
downstream. In the initial pollution history of a tidal river, 
this would would be more likely than later on when the accumula¬ 
tive pollution may build toward a uniform pollutant concentration. 

b. Uniform recirculation model . The Chester River, as a 
tributary estuary, is moderately stratified into two layers as 
described by Pritchard (1967). The net advective surface water 
flow is in the upstream direction while that in deeper water is 
downstream. Suspended or dissolved matter is dispersed in both 
directions as a result of two processes: (1) diffusion of the 
dissolved solute across the thermocline so that it is spread into 
layers whose net motions are in opposite directions. It is likely 
that the nonpolar plasticizers will quickly (2) sorb onto su¬ 
spended sediment and then settle down to the leptopellic layer on 
the sediment surface, and then either deposit or become resuspend¬ 
ed into a circulation cycle. These circula-processes may cause 
wide mixing, motably upstream as well as downstream. 

c. "Hot spots." As noted in Han's dye tracer experiments 
(1972), a pollution plume is apt to remain intact during the first 
few tidal cycles. That pattern was shown to exhibit selective 
localization effects. This would produce the opportunity for hot 
spots where an elebated concentration of pollutant may deviate 
significantly from the model based on complete mixing. 

PROBLEM STATEMENT 

It seems fundamental that environmental influx of toxic 
pollutants is feasible for study when the pollutant is identified, 
or at least limited to be among a group of candidates. In the 
present study, this line of first approach was blocked from the 
beginning because none of the affected oyster tissue had been 
saved. The circumstances are such that the toxic agent (if there 
was any) may have been introduced, reached toxic levels, and re¬ 
turned afterwards to pretoxic conditions. Although alkyl 


43 





phthalates are readily degraded by bacteria, photolysis, and 
hydrolysis, it is our belief that the known persistence of alkyl 
phthalates in the marine environment is the result of rapid sorp¬ 
tion by suspended clay-rich matter which eventually settles 
through sedimentation. Once sorbed, the rate of degradation is 
indefinite, but it is slow enough that the phthalates are con¬ 
sidered "persistent." In view of the rapid bioturbation process, 
surficial sediments (upper 10 cm) are quite likely to carry 
memory effects due to accumulative pollution during the previous 
decade. 

The difficult part in the present study was to develop 
reliable methods of measurement since it was felt that none of 
the initially available measurement technology was reliable 
enough or accurate enough for the purposes of this study. 

The principal focus in the work to be presented next is 
based on the significance of the clay-rich sediment regions be¬ 
low a 2-8 m water column. These should contain the residual 
compounds that may help to understand whether threatening or 
tpxoc plasticizer concentrations exist now, or were likely within 
the past decade. Since sedimentation occurs at an estimated rate 
of 1 cm per year, our efforts were focused on values in the top 
10 cm. 

Based on preliminary studies we initially picked sampling 
sites included apex of the mortality. The sites were chosen as 
likely to show whether or not the alkyl phthalates could have 
caused the oyster mortality. The first region is Tenneco Pond 
where the sediment composition was found to be highly polluted. 
The second region involves a series of five samples taken at the 
apex of the oyster mortality. That is where the downstream 
mortality movement was not clearly in evidence. The third region 
is at the mouth of the Chester River. That region is likely to 
contain a representative mixture of pollutants from the upper 
Chesapeake Bay. Since this region is a reference point for pos¬ 
sible upstream river pollution, we used a homogenized sample of 
the river mouth sediment "R" for reference purposes and for in¬ 
dependent analysis. 

Efforts were also made toward development of methods for 
analysis of oyster tissue. We underestimated the problems here 
so that aspect of the study was the last to be completed. These 
results will be presented later in this report. 

EXPERIMENTAL PROCEDURES 

Trace Analysis Techniques 

The key goals in trace organic analysis of sediment are to 
gchieve complete extraction, to avoid loss, to prevent contami 
and to avoid interferences. In preliminary experiments we found 


44 



that ultrasonic extraction of dry Tenneco Pond sediment with 
methanol gave easily measured quantities of DOP. Later, tetra- 
hydrofuran was found to give a higher yield. Then, a series of 
extracting solvents was tested comparatively and dichloromethane 
was found to give the highest extractive efficiency and relative¬ 
ly short times were required. The duration of the ultrasonic 
agitation was varied until it became certain that longer extrac¬ 
tion periods did not give greater extractive yield. 


The true extractive efficiency is not subject to direct 
measurement. Spiked addition of analyte produces a different 
sorbed state than the analyte in the sample matrix. To assess 
the apparent accuracy, two samples were selected for independent 
measurement as will be described. The approach was to do what 
could be done to show that re-extraction by baried methods did 
not lead to a higher yield. 

A major problem of contamination arises when measuring 
traces of plasticizers. The environment is widely contaminated 
by these substances, and the chemical laboratory is no exception. 
Their presence has been reported in bottle cap liners, solvents, 
extraction thimbles, preconcentrating resins and adsorbents, 
aluminum foil, glass, wood, air and pipet fillers (de Zeeuw 
1975, Singmaster et al. 1976, Webster and Nickless 1976). 

It is obvious that the reduced contamination can be achieved 
by deliberately minimizing the overall exposure to the sources 
of contamination by minimizing solvent volume, the number of 
manipulations, avoiding contact with any plastic materials, and 
ultracleaning of apparatus. This can be summarized in two 
statements: the sampling and processing apparatus must be un¬ 

contaminated, and the amount of methodology must be kept to an 
essential minimum. 

Although these aspects led us away from available methodo¬ 
logy (Giam et al. 1976), the end of the study showed that our 
method did come close to similar techniques by Dr. William 
Budde (EPA-Cincinnati). The procedure used by Budde (ultra- 
sonication, GCMS-selected ion monitoring) differs mainly in the 
technique used for drying the sample--direct addition of Na 2 SC >4 
to partly dried sediment. Our method uses the same desiccant 
but the drying efficiency is greater. Regardless, the two ap¬ 
proaches are appropriate for interlaboratory comparison. 

The measurement approach is based on a reported (Watson 1976) 
application of GCMS where the MS section is used in the selected 
ion monitoring mode (SIM). This provides an impressive gain in 
selectivity since only the predominant fragment ion masses are 
monitored. As a result there is a corresponding boost in 
instrumental sensitivity. The technique uses instrumental de¬ 
tection selectivity to replace the lesser certainty of multiple 
extractive and separation conditions (Giam et al. 1976). 


45 


The basis for the analytical methodology can be seen in the 
following way. The upper curve in Figure 7 is a gas chromatogram 
of a dichloromethane extract of a dried sediment sample from the 
Chester River mouth. This is the working standard, sample "R," 
located in Figure 5. The omission of any cleanup procedure 
creates an unresolved bunching of chromatographic peaks due to 
the many thousands of compounds that are present in the extract. 
However, when the mass spectrometer is used as GC analyzer, an 
extraordinary boost to the selectivity of the combined instru¬ 
mentation is realized. This is shown for mass 149 as the lower 
curve in Figure 7. The integration of m/z 149 intensity to 
measure alkyl phthalates was confirmed by comparing integrated 
area ratios for confirmational fragment ion masses for DBP 
(m/z 2-5,223) and for DEHP (m/z 167,279). This was done for con- 
firmational purposes to show that no interferences were present 
at the levels being investigated. 


It has been concluded that there is no ambiguity in the pre¬ 
sent dual use of multiple ion monitoring to identify and measure 
DBP and DEHP in a single GCMS-SEM experiment. This is a result 
of the extreme molecular selectivity, and correspondingly justi¬ 
fied methodological simplication. The generic term DOP (dioctyl- 
phthalate) will be used to refer to one or both of the isomers, 
DNOP (di-n-octylphthalate) and DEHP (di-2-ethylhexylphthalte). 
Distinction between these substances was obtained later in the 
study (see Figure 14). 


Total Ion Current 



Time (minutes) 


Figure 7. GCMS of Chester River mouth sediment extract 


46 











Cleaning Procedure 


Scrupulous cleaning procedures were used to minimize organic 
contamination of the material used in this work. The details 
are given in the following. 

Glassware (Sample bottles, vials, lab glassware, all 
glass materials ) 

1. Wash with Aquanox detergent. 

2. Rinse with tap water followed by filtered, 
distilled water. 

3. Bring to annealing temperature of the glass 
(450°C) for at least 1 hour. 

4. Slowly cool (>4 hours). 

5. Cover exposed areas with baked aluminum foil. 

6. Cover with clean caps or lids. 

Tools (Forceps, trowels, spatulas, etc.) 

1. Clean with Aquanox. 

2. Rinse with charcoal filtered water. 

3. Dry in air. 

4. Wrap in aluminum foil. 

Piston Core Liner (Brass) 


1. Scrub with Aquanox detergent using brush 
fixed on ram-rod. 

2. Rinse with tap water. 

3. Add chromic acid/sulfuric acid cleaning 
solution to etch surfaces: tube is in¬ 
verted several times. 

4. Rinse with tap water. 

5. Inspect for bright shiny inner surface. 

6. In field, tube was rinsed by brush and 
rinsed between cores using the on-board 
water supply. 


47 







Bottle cap liners (Teflon) 


1. These liners were cut from Teflon sheet using 
cork bores. They were then extracted for 

24 hours using Soxhlet extraction. 

2. Liners were stored in an air-tight, cleaned 
glass bottle with an aluminum foil liner. 


Solvents 


1. Water was purified by passing in succession 
through activated carbon filter, XAD-2 column 
(Chesler et al. 1976) and final ultrafilter. 

2. All other solvents—dichloromethane, MeOH, 
etc.—were distilled in all-glass distilla¬ 
tion apparatus. Tetrahydrofuran was di¬ 
stilled from freshly cut sodium, an important 
recommended procedure that prevents buildup 
of explosive peroxides. 

Septa (GC-Injection Port) 

1. Septa with high thermal stability were used: 
Thermogreen (TM) LB-1 Septum, SUPELCO, INC. 

2. The septa were stored in capped organic-free 
containers until use. They were inserted 
into the injection port using cleaned tools. 

This procedure made it unnecessary to precondition the 
septa after installation in the GC or GCMS. 

Field Sampling Procedures 


Water Extraction using Sep Pak Cartridge 


1. Luer-type syringe was fitted with a C13 Sep Pak 
(Waters Associates). 

2. A clean glass jar was immersed to obtain a 
subsurface water sample. 

3. This water was transferred into the barrel of 
the syringe. The plunger was inserted and the 
water was pushed through the C18 cartridge. 

4. Step 3 was repeated until the back pressure 
buildup, due to filtered particulates plugging 
the cartridge, prevented further concentration 


48 







of organics on the C18 material. The total water 
volume was noted, usually 120-150 ml. 

5. The cartridge was returned to its original foil 
packet. The foil was bent to give partial seal¬ 
ing protection. The collected samples were 
stored in a clean glass bottle that was sealed 
until analysis was ready to be performed. 

6. Trials with varied amounts of isopropanol, 
methanol and tetrahydrofuran showed that 75-80 
percent recovery was obtained following desorp¬ 
tion with at least 1.5 ml of tetrahydrofuran; 

2.5 ml was used in the actual procedure. 

7. 1.7 ml of water was added to the extract from 
(6) to match the liquid composition to the 
initial carrier composition used in liquid 
chromatography. 

Sediment— 

1. Tenneco Pond and contiguous creek sediments. 

A small grab sampler was used to take surface 
samples. The sampler was opened and the sam¬ 
ple was discharged into a scrubbed galvanized 
metal bucket. A clean metal scoop was used 
to transfer the moist sediment to clean glass 
jars. 

2. Chester River. Sampling sites were selected 
near the main channel in order to obtain clay- 
rich specimens. A Van Veen grab sampler was 
used to bring up surface sediment samples. 

The sampler was fitted with a sliding panel 
to permit insertion of the coring tube. The 
clean brass coring tube was inserted to get 
a 10 cm vertical core of the uppermost sedi¬ 
ment. The brass tube was sealed by placing 
a hand at the top. Then the cylindrical core 
was lifted out of the grab sampler. The core 
was released onto baked aluminum foil, and 
then stored in the foil in a clean glass con¬ 
tainer. The latter was stored for library 
purposes. 

3. All sediment samples were transferred within 
12 hours to storage at 4°C until ready for 
drying. 

4. All sediment samples used in this study were 
subjected to the following drying procedure. 


49 


Sediment Drying by Isopiestic Dehydration 


a. The collected sediment is homogenized in the sample 
container using a clean metal spatula. 

b. A 10 g sample is spread evently onto the surface of a 
clean 12 cm watch glass using the same spatula. 

c. An organic-free desiccator is charged with 0.25 kg 
Drierite in the desiccator at 23°C for 48 hours. 

d. To complete the drying, the sediment is removed to 
repeat the drying process. The desiccant is re¬ 
dried by baking in the desiccator at 250°C for 

12 hours. The partly dried sediment is then re¬ 
turned to the cooled freshly baked desiccant for 
another 48 hours. 

e. The dry sediment is scraped into a clean glass 
mortar. It is then pulverized using a clean glass 
pestle, transferred to labeled vials, and stored 
at 4 °C. 

Two solid substances will undergo a gas phase transfer of 
water until a final equilibrium vapor pressure is reached. The 
repeated exposure of moist sediment to repeated fresh charges 
of the desiccant Na 2 SO^ causes isopiestic dehydration. The 
advantages of this approach are as follows: 

1. Water content in clay sediment is reduced from about 
60 percent to about 1 to 2 percent on a dry weight 
basis. 

2. Unlike the procedure of mixing the desiccant in with 
the sediment (Bulla, personal communication), the 
water is physically removed and there is less oppor¬ 
tunity for organic contamination from the desiccant. 

3. Unlike lyophilization, isopiestic dehydration is 
carried out at atmospheric pressure with 10 3 -fold 
shorter gas phase collision distances and a corre¬ 
sponding less likelihood for sublimative loss of 
the measured analytes (DBP or DEHP). 

Sediment Extraction 

From Karrickhoff's work (1979), it is known that organic 
pollutants in sediment are associated with the organic carbon 
content, due principally to humic and other associated polymers. 
Based on this, the sorptive structure should be modeled effec¬ 
tively by n-octanol. Accordingly, a volative extraction solvent, 
similar to polarity to octanol, was chosen as more likely to 


50 





give efficient extraction. Since water polarity is much higher 
than that of octanol, the partitioning model clearly infers the 
need to remove water before attempting the extraction of neutral 
organic compounds. 

This approach differs considerably from the conventional 
Soxhlet extraction procedure applied to wet sediment. The lat¬ 
ter also requires large solvent to sample ratios so an extra 
burden is implied in terms of needed solvent purity. We were 
advised by Dr. George Boughman (personal communication) that 
Soxhlet extraction tends to be incomplete due to slow diffusion 
of solvent through the sediment sample. For that reason, a thin 
coating of sediment is advised to line the extraction thimble. 
Unfortunately, this procedure boosts the solvent-to-sample ratio 
even further. 

Sediment Extraction with Methylene Chloride 


1. Two grams of sediment is weighed to the nearest mg 
and placed in 10 ml capacity vials with Teflon 
lined caps. The caps are screwed on tightly to 
prevent evaporation. 

2. Five ml of methylene chloride, containing 0.40 ppm 
dimethoxyethyl phthalate (DMEP) as internal standard, 
is introduced using a Repipet apparatus. (A different 
amount of DMEP was used for the Tenneco Pong sample. 
This involved 4.0 ppm DMEP in CE^Cl^, but there was no 
subsequent evaporation.) 

3. The vials are first agitated in a Vari-whirl mixer to 
suspend the particles. 

4. The vials are placed in an ultrasonic bath. Circu¬ 
lating water is used to hold a temperature of ap¬ 
proximately 30°C for 2 minutes. 

5. The vials are centrifuged at 2,500 G for 15 min. 

This is done to remove suspended particles. 

6. The supernatant liquid is drawn off and placed in 
a 5 ml capacity vial using a Pasteur pipet. 

7. The extract is evaporated under a stream of purified 
nitrogen to a volume of approximately 0.5 ml. The 
nitrogen is purified by passing through a column of 
carefully extracted washed (Chester et al. 1976) 

XAD-2 resin. 


51 



8. Two hundred pi of isooctane is added to the vials to 
prevent evaporation to dryness. The extract is blown 
down to 0.20 jul. This gives an overall 25 x concen¬ 
tration (5 ml to 0.2 ml). 

9. A procedural blank was prepared in triplicate for 
each experiment. Each contains the extracting 
solvent and internal standard but no sediment 
samples. Isooctane is added after the blanks 
have been sonicated and blown down. For LC 
analysis, tetrahydrofuran was substituted for 
the isooctane. These procedures are summarized 
in Tables 9 and 10. 


TABLE 9. FIELD SAMPLE HANDLING 



Procedure 

Precleaning 

Surface Grab 

Sampler 

On-board 

High Pressure 

Water Hose 

Obtain Core within 

Grab Sampler 

Use brass tube precleaned 
by acid etch, soap, water. 
Swabbed with abrasive 
between samples. 

Storage 

Use baked glass bottles, 
aluminum cap liners. 

Hold for 14 days at 4°C. 

Isopiestic 

Dehydration 

Two 48-hour periods. 

Closed system 

Sample 

Homogeneity 

Sample grinding with 
baked glass mortar and 
pestle. 

Storage 

Dried sample stored at 

4 °C. 


52 







TABLE 10. METHODOLOGY 



Step 

Procedure 

Reagents Added 

Conditions 

1 

Isopiestic dehydration 

Na^O^dried 

Closed system 


of sediment 

at 180 °C 

22°C, 1 atm 

2 

Ultrasonic extraction 

CH 2 CI 2 contain- 

Closed system 


with CH~C1 0 

ing 0.4 or 4.0 

t = 30 °C 



ppm DMEP internal 



standard 


3 

Centrifugation 

None 

Closed system 


t = 10°C 
2500 x G 


4 Evaporate solvent XAD-2 filtered Open system 

N 2 : 

(a) Isooctane 
added for 
GC or GCMS 

(b) THF added, 

CH 2 C1 2 re¬ 
moved, for 
HPLC 


Preliminary Liquid Chromatographic Analysis 


High performance liquid chromatography has been reported 
for analysis of alkyl phthalate esters (Heilman 1978) . This 
technique was adapted to confirm GC measurements on Tenneco 
Pond sediment, as a screen and to provide upper limit data on 
samples containing small amounts of alkyl phthalates. The 
Heilman method is based on adsorption chromatography. A more 
reliable method was developed based on partition chromatography 
using water-tetrahydrofuran as the carrier with bonded C18 as 
the stationary phase. 

A surface sample of water from the environment was trans¬ 
ferred onsite to the barrel of a baked glass Luer-type syringe. 

A Sep Pak C18 cartridge was already fixed in place. The water 
was then passed through the cartridge using hand pressure after 
inserting the plunger into the barrel of the syringe. This was 
continued until the back pressure became too great. Then the 
cartridge was returned to its original protective envelope, 
sealed, and labeled. 

Preliminary laboratory tests had shown that THF (tetrahydro- 
furan) gave sharper peaks than either methanol or isopropanol 
which are widely used. The recovery of DEHP standard was about 


53 








74 percent by desorbing with 1 ml of THF, or 75-80 percent if 
> 1.5 ml were used. The final procedure called for 2.5 ml of 
THF. Then 1.7 ml of purified water was added so that the 
liquid sample composition matched the initial HPLC carrier 
make up. The liquid chromatograph 254 nm UV monitor (Waters 
Model 440) remained on scale using 25 pi samples of extract. 

Similar tests were carried out using DTDP (ditridecyl- 
phthalate) and the results were quite similar except that a 
lower recovery, approximately 60 percent, was obtained. 

The precision of the HPLC results were found to be linear 
with the amount of added standard to within 5 percent variabil¬ 
ity. This was observed during Sep Pak adsorption of 10 ppm 
DEHP dissolved in water samples ranging from 30 to 150 ml in 
volume. 

Water analysis in Tenneco Pond and several sampling sites in 
the Chester River was attempted using C18-Sep Pak cartridges to 
concentrate the analyte. These cartridges were soon clogged by 
suspended particles so the sample volume was limited to 150 ml 
or less. GCMS showed that the liquid chromatographic (LC) 
technique alone in one instance gave an apparent but false 
identification of ditridecylphthalate. At trace concentrations, 
LC was considered valid for setting upper limits. 

The final conditions for HPLC analysis were obtained using 
a linear carrier gradient proceeding from 60 percent THF/40 
percent H 2 O to 90 percent/10 percent over a 5.0 minute period. 

A flow rate of 2.0 ml/min was used. 

Sediment extracts in dichloromethane were twice treated by 
adding THF and using purified nitrogen gas blowdown to remove 
the dichloromethane. Then makeup water was added, as before, 
to give the correct liquid solvent rates. 

Quantitative Analysis of Sediment Extract using GC/MS-SIM 

The Hewlett Packard 5992A GC/MS system was used for the 
analysis of sediment extracts. The microprocessor-controlled 
5992A uses a jet separator to interface the gas chromatograph 
to a hyperbolic quadrupole mass filter. A 3-foot (90 cm) 
silanized glass column (packed with 3 percent SE-30 on Chromo- 
sorb W-AW-DMCS) was used. The column was temperature program¬ 
med from 140°C-250°C at 5°/min. The system was later converted 
to a SE-52 glass capillary column programmed from 160°C-275°C 
at 7.5°/min. Up to 6 ions can be monitoring during a chromato¬ 
graphic run in the selected ion monitoring mode. The base peak 
of DEHP, m/z 149 was monitored. Mass 59 was also monitored 
which is characteristic of the internal standard, dimethoxyethyl 
phthalate (DMEP). DMEP has a low mass 149 abundance which de¬ 
creases its chromatographic interference with other possible 


54 



phthalates. It is also likely to exhibit physical and chemical 
properties similar to other phthalates, and it is not produced 
commercially as a plasticizer. By adding DMEP to the extracting 
solvent, methylene chloride, it is susceptible to the same sys¬ 
tematic errors during the analytical scheme as other phthalates 
being determined. 

GC/MS Autotune Procedure 


Each week "autotune" was run to adjust and to assess the 
condition of the GC/MS instrument. This procedure tunes the 
ion source and mass filter to produce a mass spectrum of per- 
fluorotributyl amine (PFTBA) to meet certain minimum specifica¬ 
tions. If these specifications, as recommended by the 
manufacturer, were not met, the problem was diagnosed and 
corrected before continuing the work. 

GC/MS-SIM Calculation 


The mass 149 chromatogram of all sediments taken from the 
Chester River show only two well-defined peaks at GC retention 
times that correspond to dibutylphthalate DBP, and dioctyl- 
phthalate. Quantitative analysis for each of these two 
compounds is based on monitoring mass 148 peak areas for DBP and 
DEHP, and mass 59 for the internal standard. Within experimen¬ 
tal error, we verified the linearity typical in GC/MS-SIM 
analysis. Defining the following terms 

C = concentration of solute in CI^C^ (mg/ml CI^C^) 

S - area under mass-chromatographic peak (area units) 
a = analyte (DEHP or DBP) 

i - internal standard (DMEP) 
k = response factor 


we have 


S = K and S. = K. c.. 
a a ill 


The ratio k a /k. was found to be constant for total analyte in¬ 
jections less than 0.3 pg providing C /C. were within a factor 
of 100 of unity. Then, it follows that * 1 


R = 



constant 



C 

a 




C 


a 


S. 


l 


55 






The latter term was measured giving k /k. = 11.0 + 0.4. It 
follows that a i — 


In order to calculate the weight fraction f of analyte in 
the sediment sample, the following relationship a was used: 


f 


a 


. jig of a 
g sediment' 




of a 


CH 2 C1 2 


) 


V e (ml CH 2 C1 2 ) 
W (g sediment) 

o 


This uses the volume of the final concentrate extract, and 
the initial weight W g of dry sediment to provide the weight of 
DBP or DEHP. The units, pg/g, are the same as parts per million 
(ppm). 


RESULTS AND DISCUSSION 
Preliminary Results with HPLC 

A detailed study (Gingras 1979) was made of the basis for 
applying modern liquid chromatography to analysis of field 
sediment and water samples for alkyl phthalates. Previously, 
Heilman (1978) had developed the use of adsorption liquid chro¬ 
matography but this is prone to irreversible adsorption effects 
and lesser reproducibility. Amundson (1978) used reversed phase 
LC with Bondapak C-18 as the stationary phase with methanol plus 
1 percent acetic acid as the carrier. More sharply defined 
peaks were obtained more rapidly with tetrahydrofuran-water 
mixtures, 60:40 linear programmed in a 5-minute period to 90:10 
at 2.0 cm^/ml flow rate. 

Analysis was made by LC bases on preliminary test samples of 
phthalates as manufactured, named and provided to us by Tenneco. 
These are shown in Figures 8-12. The LC technique has notable 
stability but qualitative identifications can only be shown to 
be consistent or inconsistent with the LC retention volumes. The 
time of analysis was only 7 minutes, or less than half required 
by Anumdson. The resolution of a synthetic mixture of the three 
reference materials DOP, DIDP (diieodecylphthalte) and DTDP is 
illustrated in Figure 9. Later on in the study we established 
that Tenneco's "DOP" was actually DEHP. 

Analysis of Tenneco Pond water was carried out using the Sep 
Pak technique and the results are shown in Figure 10. The pre¬ 
sence of DOP 0.25 + 0.15 ppm and DTDP (ditridecylphthalate, 

1.5 + 0.2 ppm) were estimated by LC alone. The presence of 
phthalate esters in downstream water samples could not be de 
tected by LC so these will not be reported here. The first peak 


56 






UV Detector Response 



Time (minutes) 


Figure 8. Liquid chromatography of Tenneco Products de¬ 
signated by Tenneco as Alkyl Phthalates, or mixtures (6-10P 
and 7-11P). 


57 





































UV Detector Response 



Figure 9. Liquid chromatogram of DOP, DIDP, and DTDP 

(synthetic mixture of Tenneco samples). 


5 8 











UV Detector Response 



Time (minutes) 


Figure 10. 


Liquid chromatogram of Tenneco pond water. 
Identification based on Tenneco reference samples. 


59 
















in Figure 10 is likely to be due to the fulvic acid component 
which occurs in natural water and sediment extracts. Although a 
number of Morgan Creek and Chester River water samples were run 
by the same technique, the method was frustrated by early plug¬ 
ging of the Sep Paks. This prevented the obtaining of large 
enough samples to give the needed sensitivity. 

Analysis was made by LC of Tenneco Pond sediment. The re¬ 
sult is shown in Figure 11. The concentrations are so high that 
peak verification by GC/MS was readily demonstrated and quanti¬ 
tative interpretations are thus possible. The retention volumes 
are consistent with the presence of humic acid, of DEHP 
(1.8 + 0.1) x 10 3 ppm and a second determination of (1.3 x + 0.2) 
x 10 3 ppm, of DIDP (1.4 +0.2) x 10 3 ppm and of DTDP 
(1.9 + 0.2) x 10 3 ppm. The qualitative identifications by LC 
were each confirmed by GC/MS-SIM retention times of m/z 149. 

A few miles downstream from Tenneco on Morgan Creek sediment 
samples were taken for LC analysis. Comparison was made to 
humic acid under the same conditions. These results are shown 
in Figures 12 and 13, respectively. It is clear that the natural 
background, the multiple peaks due to fulvic and humic acid con¬ 
stituents, seriously interfere with and prevent quantitative use 
of these LC results. Sediment samples taken from the 
Chestertown Bridge and from the Chester River (site 5, Figure 4) 
showed no evidence for DEHP, apparently less than 1.0 ppm. 
Artifact peaks were observed and these suggested exaggerated 
levels of DIDP and DTDP. The GC/MS technique gave qualitative 
but not quantitative confirmations of DEHP and DBP so these were 
concluded as present at both sites. The DBP peak position was 
found to be consistently obscured in the LC by the humic com¬ 
ponents. Significantly, this was the only alkyl phthalate which 
Tenneco did not make, but which is otherwise massively produced 
by U.S. industry. Second, certain peaks evident at the equivalent 
of <10 ppm levels by LC were found by GC/MS not to be phthalates. 
Attention was then directed to the GC/MS technique which was con¬ 
sidered necessary for identification and measurement of DBP and 
DEHP at levels below 100 ppm. The LC technique offered prelimi¬ 
nary qualitative utility and it was the more reliable for meas¬ 
uring DIDP and DTDP. 

Preliminary Qualitative Results Using GC and GC/MS 

A preliminary grab sample of Tenneco Pond sediment was 
obtained. The sediment was desiccated and then extracted by 
methanol using ultrasonic agitation. Centrifugation gave a 
clear filtrate which was then analyzed by gas chromatography. 

The result showed a single peak which accounted for >95 percent 
of all volatiles apparent by the FID detector. 

Gas chromatographic retention times and mass fragmentation 
patterns showed clearly that the very conspicuous main organic 
volatile component in Tenneco Pond sediment is the same as the 


60 



DEHP reference sample which, in turn, matches the samples of DEHP 
from Chem Service, Inc. The mass fragmentation patterns showed 
closely similar abundance profiles at m/z 61, 70, 71, 83, 104, 
112, 113, 149 (base), 150, 167, 168, and 279. 



Figure 11. Liquid chromatogram of Tenneco pond sediment. 


61 






Reference data from the NIH/EPA library, recently published by 
the National Bureau of Standards, strengthened the assignment 
of DEHP. 

DEHP was subsequently distinguished from DNOP by their g.c. 
retention times and mass spectra. There was no DNOP detected in 
the Chester River sediment. The use of glass capillary GCMS 
permitted a clear distinction (see p. 69) . 


Tests of the Methodology Based on GC/MS 

A series of experiments were carried out in order to 
validate the method. Preliminary experiments showed that de¬ 
sorption of DEHP or DBP occurred more efficiently from dry 
rather than wet sediment. Very few wet extractions were 
carried out thereafter. 


Rate of Extraction— 

A series of extractions by dichloromethane of the dry work¬ 
ing standard "R" (see Figure 7) were carried out. The DBP 
extraction was independent of the time given to the sonication 
step which was varied from 30 seconds to 4 hours. The results 
are shown in Table 11. The results show no apparent trend with 
time of sonication. Other experiments confirmed the finding 


TABLE 11. EFFECT OF SONICATION ON DBP EXTRACTION BY CE y Cl ? 

FROM SAMPLE "R" Z Z 



Sonication 
Time (min) 

> 

DBP Measurement 
(ppm)* 

0.5 



0.785 * 

5.0 


0 . 

725, 0.66 

30.0 



0.75 

120.0 


0 . 

79, 0.715 


Average Value 

0.74+0.02 (+ SDM, n = 6) 


Standard 

Deviation 

0.05 


Standard Deviation of Mean 

0.02 


Internal 

Standard: 

d-10-Anthracene 


These preliminary results are to be used on the basis of their 
relative accuracy. Subsequent determinations of absolute DBP 
levels were found to be more accurate. 


62 










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that the desorption rate in CH 2 CI 2 was too rapid to be observed 
kinetically. The extraction time was fixed at 2.0 minutes. It 
should be noted here that reproducible time course studies were 
not demonstrated until we discovered the necessity of using 
cooling water (30°C) to prevent a temperature rise and sub¬ 
sequent loss problems during the extraction period. 

Effect of Solvent— 

Various solvents were tested for their extractive power. 

The conditions were identical to those used in Table 12. The 
results are shown in Table 12. It is clear that hexane gives 
decidedly poor extractive efficiency, while benzene, dichloro- 
methane and methanol give quite similar extractive efficiency. 

The choice of solvent depends upon more than the rate of 
extraction. The variation in total amounts of extracted organic 
matter was not measured. A simple color comparison of extracts, 
often clarification by centrifugation, showed that hexane exhi¬ 
bited least color, dichloromethane gave a visible light yellow 
color, and methanol showed a substantially deeper color. 

Analyses were performed on the extracts immediately after ex¬ 
traction. After a period of several weeks a qum would form in 
the dichloromethane extract and inconclusive tests suggested 
that dissolved organics might have been lowered in concentration 
as a result. Clearly, methanol would lend to a distinctly 
higher concentration of nonvolatile substances and these would 
be likely to foul the chromatograph. Hexane gave the cleanest 
extract but, of course, it has a poor yield. Benzene is toxic 
so this was ruled out. Therefore, dichloromethane was chosen 
for its high extractability, low toxicity, and ease of 
volatilization. 

TABLE 12. COMPARISON OF ULTRASONIC EXTRACTION EFFICIENCIES 


OF VARIED SOLVENTS MEASURED ON WORKING STANDARD* 



Solvent 

Trials 

Measured DBP 
(ppm) 

Hexane 

3 

0.34 + 0.07 t 

Benzene 

3 

0.65 + 0.02 

Dichloromethane 

3 

0.70 + 0.01 

Methanol 

3 

0.67 + 0.03 


Based on a fixed ultrasonic extraction period of 2.0 minutes 
(Chester River Sample "R"). 

Tolerance is expressed as the standard deviation. 

Internal Standard: D-10 anthracene. 


64 







Recovery Tests— 

Several experiments were carried out to show whether DEHP 
could be added to the measurement system and then recovered 
without serious loss. The results are summarized in Table 13 f 
which is largely self-explanatory. The most serious loss seemed 
to occur when dichloromethane containing a DEHP spike was de¬ 
liberately evaported to remove all of the solvent. When this 
was done a 7 percent DEHP loss was observed. To prevent this 
in the actual procedure, a small amount of isooctane, a higher 
boiling solvent, was added to prevent the volatilization and 
no other measurements in the report involved total solvent 
volatilization exhibited here for test purposes. 


TABLE 13. DEHP RECOVERY MEASUREMENTS USING DICHLOROMETHANE 



Sample 

Procedure 

Trials 

Percent 

Recovery 


Spiked Solvent* Remove 95% of solvent. ^ 

Add fresh solvent to 
bring to initial volume. 

Spiked Solvent* Remove 100% of solvent. 3 

Add fresh solvent to 
bring to initial volume. 


100 + 2 

93 + 1 


Spiked Blanki 
(attapulgite) 


Remove 98% of water by 4 

desiccation. 


Spiked Blank 
(attapulgite) 


Ultrasonic extration by 
dichloromethane 


98 + 1 


5 ml of CH 2 C1 2 containing 1 mg of added DEHP. 

1 g of Attapulgite containing 0.2 mg of added DEHP. 

N.B.(1) Internal standard = d-10-anthracene 
N.B. Reagent blanks are as follows: 

CH 2 C1 2 = 20+10 ppb of DEHP 

Attapulgite = 10 ppb of DEHP 


Effect of Varied Methodology and Combinations— 

The establishment of a 2.0 minute sonication procedure for 
extracting dry sediment with dichloromethane was subjected to 
a series of tests to measure the completeness of the extractive 
process. The same working standard "R" was used. The sonica 
tion procedure is the same as that previously described. The 


65 








Soxhlet extraction procedures are as follows: 


Wet Soxhlet Procedure 


1. The extraction thimble 43 x 123 mm was extracted 
with methylene chloride for 24 hours before use. 

2. A clean 2000 ml flask was charged with 500 ml of 
methanol containing approximately 5 pg of the 
internal standard, d-10 anthracene ( C ]_ 4 D io) * 

3. The thimble was loaded with approximately 90 g 
of wet sediment (60 percent moisture content). 

The sediment was smeared onto the walls of the 
thimble cylinder to provide more intimate con¬ 
tact with the solvent. 

4. After 24 hours the methanol was emptied into a 
storage container and replaced by 500 ml methylene 
chloride and an additional 5 pg of d-10 anthracene. 

5. The extraction was continued for at least 48 hours, 
when the extract solution in the thimble chamber 
became completely clear. 

6. The solvents were distilled off using Buchi Roto- 
vapor R. until approximately 30 ml of aqueous 
extract remained. 

7. This aqueous liquid was extracted with 5 volumes 
of methylene chloride. This extract was concen¬ 
trated to 5 ml using the Rotovap. Further 
evaporation was done using a stream of nitrogen 
(XAD-2 filtered). 

8. A blank was prepared by duplicating the procedure 
described above, but without the sediment samples. 

Dry Soxhlet Extraction —This procedure is the same as the 
wet Soxhlet extraction procedure except for the following modi¬ 
fications . 

1. The sediment was first dried by desiccation to a 
moisture content of 1-2 percent. 

2. Approximately 40 g of dried sediment was added to 
the previously extracted thimble. 

3. Only methylene chloride was used as extracting 
solvent and the extraction duration was 72 hours. 


66 




4. 


The methylene chloride extract was concentrated 
directly to 5 ml using the Rotovap followed by a 
stream of nitrogen. No solvent extraction step 
was needed. 

The results of the various extraction tests are as 

follows: 


Test 1 

A comparison of the efficiency of ultrasonication 
and Soxhlet extraction is shown in Table 14. Ultrasonication 
extracted greater amounts of phthalates than Soxhlet extraction 
from the same sediment collected at the mouth of the Chester 
River. The incompleteness of the Soxhlet method is believed to 
be the result of the lower eddy diffusion of solvent in the com¬ 
pacted solid sediment. With particular regard to the time course 
study, it is felt that the present experiments give a sound 
basis to reject the Soxhlet technique on the basis of its 
greater proneness to contamination, its incompleteness and the 
high cost associated with slow rate and need for laborious re¬ 
petition . 


Test 2_ 

A second and more stringent test of the sonication 
technique was carried out. Sonicated material was rinsed free 
of retained extracting liquid, and the samples were then 
subjected to re-extraction. The results are shown in Table 15. 

The repeated use in test 26 of sonication drew a 
blank—no evidence for further extraction was observed. Es¬ 
sentially the same finding was observed when the sonicated 
sample was washed free and then subjected to Soxhlet extraction. 

Test 3 

A comparison was made of wet Soxhlet and dry sonica- 
tive extraction of Tenneco Pond sediment. In general, these 
more heavily polluted samples of sediment seemed to be more 
easily extracted so that is not a stringent test. 

Analysis of the Internal Standards --The use of dimethoxyethyl 
phthalate (DMEP) seemed ideal in the sense that its gas chromato¬ 
graphic elution time fit into a window that caused no inter¬ 
ference with the DBP or DEHP measurements. Further, its 
chemistry was parallel to that of the other phthalate esters. 
However, we eventually became aware that our procedural blank 
levels were significant: 0.10 + 0.04 (n=6) ppm for DBP and 
0.28 + 0.22 (n=3) ppm for DEHP. Direct analysis of the DMEP 
standard revealed the cause since it contained 4 percent DBP and 
11 percent DEHP! These are not serious interferences since the 


67 






TABLE 14. COMPARISON OF METHODS FOR EXTRACTION OF PHTHALATE 
_ ESTERS FROM CHESTER RIVER, MOUTH SEDIMENT. TEST 1. 

Extraction Method Amount Extracted, ppm , and 

Standard Deviation 

/ DEP / DBP / DEHP 

Ultrasonication 0.19 + 0.03 0.36 + 0.07 0.40 + 0.06 

Soxhlet, dry 0.10 + 0.03 0.33 + 0.14 0.34 + 0.06 

Soxhlet, wet 0.05 + 0.10 0.26 + 0.07 0.21 + 0.12 

Re-extraction by ultrasonication of sediment from all three 
methods yielded less than 1 percent of the first extraction value 
for all three phthalates. 


\ 


TABLE 15. TEST OF VARIED EXTRACTION AND RE-EXTRACTION PROCEDURES 



Sample 

Test 

Method 

Sediment Trials Result (ppm) 
State 


Chester 

River 

1 

Soxhlet and 

Dry and 

3 

See Table 

Sample 

"R" 


sonication 

wet 



(same) 


2a 

Soxhelt 

Dry 

3 

0.9+0.1(DBP) 




Re-Sonication 

II 

3 

0.04+0.02(DBP) 

(same) 


2b 

Soxhlet 

II 

4 

0.7+0.1 (DBP) 




Re-Sonication 

II 

3 

0.02 (DBP)t 




Re-Soxhlet 

II 

1 

0.02 (DBP)f 

Tenneco 

Pond 

3 

Soxhlet 

Wet 

1 

1.0xl0 3 (DEHP) 




Sonication 

Dry 

5 

(1.2+0.1)xlO 3 


(DEHP) 


+ Uncertainties are expressed as standard deviations of the mean 
values (rounded up). 

^ i.e., none detected. 

Internal Standard: D-10-anthracene 


68 












calculations used a conventional blank subtraction step. How¬ 
ever, the use of the contaminated DMEP was not considered to be 
desirable and another standard was introduced. 

Since the method verification and environmental measurements 
were made at different times, we have been careful to note the 
internal standard for each experimental series. 

The more recent work involved the use of an excellent in¬ 
ternal standard, D-10-anthracene, C^D^o* A small amount of DBP 
and a negligible amount of DEHP were observed with the anthracene 
standard. A correction for the DBP in the average blank was used 
in the caluculations. The use of C^D-^q standard allowed more 
reliable measurements in the sub-ppm region. The basis for these 
statements, for the GCMS capability for discriminating between 
DEHP and DNOP, for the virtual absence of DNOP from sediment, and 
for the purity of the D-10-anthracene internal standard, i.e. 
illustrated in Figure 14. 

Interlaboratory Comparisons of Split Sediment Samples 


Two samples were carefully homogenized and sent to Dr. 
William Budde (EPA - Cincinnati) for interlaboratory comparison. 
The samples are the Chester River working standard "R" and a 
Tenneco Pond sample. The results are shown in Table 16, A much 
closer agreement exists between the Tenneco Pond than the poor 
results obtained on the initial "R" sample. Since this involved 
two essentially independent experiments on the two unrelated 
samples and standards, it was decided to accept the results as 
validative for Tenneco Pond and to reject the results on sample 
"R" as nonvalidative. 

The interlaboratory comparison was repeated using a fresh 
sample from our working standard "R." Samples were sent with and 
without use of our drying procedure. In addition, a spiked blank 
sample containing 200 ppm DEHP was sent for comparison measure¬ 
ments. Measurements on the split portions that were saved for 
that purpose were repeated. The results are given in Table 16. 
The results show reasonable agreement between the two labora¬ 
tories. It should be noted that the results from EPA- 
Cincinnati seem to be higher than our results, and this remaining 
problem is still under study. However, the main purpose of 
demonstrating comparability has been achieved. A chronological 
account of interactions with Dr. William Budde (EPA- 
Cincinnati) is presented in Appendix B. 


69 



A. mixture of known standards (Analabs, Inc.) 
iVc * 149 



C. 3 viL injection from 200 uL total extract 
from 5.0 g of dry Attapulgite matrix 
m/c = 149 


JL 


A 


A 


D-10-Anthracene 


\ 


D. 




main pack corresponds to 6.0 mg of internal 
standard (D-10 anthracene) 

m/c = sum of 188 (internal std.), 57, 149, 
202, 228, and 252 


i 


10 


—I_ 

15 


TIME (minutes) 


Figure 14. Comparison of GCMS-SIM chromatograms. 


70 









































TABLE 16. 

INTERLABORATORY 
SPLIT SAMPLES* 

COMPARISON OF DEHP 

MEASUREMENTS OF 


Samples 

Method 

This Laboratory 
DEHP (ppm) see 
note d 

EPA-Cincinnati 

DFHP (ppm) (n) 


Chester River, "R" 

GCMS 

0.097+0.03 (5) 

58 

see 

note 

a. 




0.3 

see 

note 

b. 




0.4 

see 

note 

c. 

Spiked Attapulgite 

GC 

200+10 (5) 

121 




(dry) 

GCMS 






Tenneco Pond West 

GC 

1200+100 (5) 

1700 





(dry) 


Notes: 

a. Initial determination. This errant result necessitated 
the subsequent repeating the entire analysis using 
freshly split samples. 

b. Dried by this Laboratory. 

c. Dried by EPA-Cincinnati. 

d. Reagent blanks measured DEP 0.0005 ppm, DBP 0.001 ppm, 
DEHP C.003 ppm. 

* 

Further details are given in Appendix B. 


Tenneco Pond and Chester River Sediments 


Sediment analyses for DBP and DEHP were measured using the 
GC (Tenneco Pond) and GCMS (Chester River) techniques. The 
results are presented in Table 17. The samples in all cases were 
surficial—top 10 cm—that were thoroughly homogenized prior to 
drying and subsampling. The results are persented along with 
percentage of organic carbon [HC1 (0.1M) treatment was used to 
remove carbonate] and percentage of water. All results are pre¬ 
sented on a dry-weight basis. 

Two samples were taken from Tenneco Pond which is roughly 
oval in shape. These separate samples were obtained near the 
center axis of the pond about one third of the distance from 
either end: these are labeled East and West. 

It is clear that the pond sediment is quite substantially 
polluted with 0.15 percent DEHP. The previously cited LC re¬ 
sults which seem to be much better for the larger alkyl phtha- 
lates show similar amounts of other compounds: 0.14 percent DIDP 


71 








and 0.19 percent DTDP. The sum is nearly 0.5 percent on a dry- 
weight basis. 

The total content of organic phthalate esters in Tenneco 
Pond can only be guessed at since the vertical concentration 
distribution was not determined in this work. For a depth of 
0.04 m, the Tenneco Pond sediment volume for 40 x 10^ m^ of sur¬ 
face area is 1.6 x 10^ m^, or roughly 6.4 x 10^ kg on a weight 
basis. The measurements are thus compatible with the presence 
of 960 kg of DEHP, 900 kg of DIDP, and 1200 kg of DTDP, or a 
total of about 3,000 kg of these three organics. 

As discussed earlier, the State discharge permit allows 
Tenneco to release total organic extractables at a rate of 
2000 kg/year. It would appear that Tenneco discharge may be 
operating rather near the permit, depending on where the 
measurement is made: plant discharge or outfall from the dam. 

The pond is clearly acting as a secondary waste treatment facil¬ 
ity, and it is obvious that less organics have been flowing out 
of the pond than those that enter. The DBP content is low be¬ 
cause Tenneco has rarely manufactured it. 

The fact that the pond levels are now high should be inter¬ 
preted as a warning. As the pond sediment becomes increasingly 
saturated with these organics, they may eventually move out into 
the Morgan Creek conduit. Since the linear flow rates are much 
higher and the creek bed is rather narrow, it is important to 
consider the eventual saturation of the present sorptive capacity 
of the pond sediment. At a future time the allowed Tenneco dis¬ 
charge, 2000 kg/year, may become more likely to make a direct 
transit from the factory site to the Chester River, and to pos¬ 
sibly high localized concentrations in the river sediments. If 
this situation is allowed to continue without any further re¬ 
straint, one can not help but visualize a more pessimistic 
future for the sediment beds in the Chester River. 

The Chester River sediments exhibit ranges of 0.020 to 0.064 
ppm for DEHP and 0.23 to 0.85 ppm for DBP. Stations 3, 4, 5 and 
6 were deliberately localized at the apex of the oyster mortality. 
The data are not marked by major apparent differences from the 
site "R" at the river mouth. Since healthy oyster beds have been 
maintained downstream of sited 4-6, there is no framework pro¬ 
vided by the present data to assign alkyl phthalates as causally 
related to the 1973-75 oyster mortality. Of course, this is 
purely circumstantial reasoning, and the results can not be used 
to rule out the possibility. However, no oyster samples were 
saved . 


TABLE 17. DETERMINATION OF SEDIMENT COMPOSITION 


to 

W 

Q 

\ 

w 

Q 


■K 

CO 

G 

O 

•H 

-P 

cd 

P 

-P 

G 

0 

U 

G 

O 

U 

0 

-P 

fO 

(0 

jC 

-p 

.c 

CU 


>i 

Pi 

iH 

< 


T3 

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rC 

-p 

0 

£ 


U 

<A° 


o 

CM 

X 

CfP 


Cn 

G 

O 

PI 

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0 

PI 


0 

-P 

•H 

CO 


G 

+- 

g 

to cu 
K d, 
W 
D 


G 

4 - 

g 

Pu Ch 
OQ Du 
Q 


G 


IP Ch 
PI D, 
Q 





o 










o 










o 



CM 







-4 


• 

• 







+ 1 


o 

O 







o 


+ 1 

+ 1 







o 


r- 

H’ 







o 


• 

• 







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o 

O 





X X 










-- . <-S 



CO 

-—V 

<—■ » 





cn co 



N —’ 

ID 

in 




CO 

• ^ • "■—^ 




'—■ 

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rH 

o 

oco oco 



• 

+ 

+ Icn 

+ 1 


1 — 

rH 

+ |o + |o 



rH 

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CO 

X CO iH CO rH 



+ l 

in o 

CM o 

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— 

CM 



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CO o 

id in 

CTl rH 

+ hr 

'— 



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rH O 

r- o 

O 

CO 

,—i 

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CM 


o • 

o • 

O • 

CM 

• 

ID 



• 


• o 

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• O 

• 

o 

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o 

o 

O 

o 


O 

















CO 


H' 






CM 

CM 

' — - 


— 

CM 





— 

—- 

+ |rH 

+ rH 

— 





r- 


ID O 

CO 

CM 

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CM 

rH 

rH O 

r- 

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o 

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o • 

o 

• 

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• 

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CO 

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CO 


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£ 

£ 

£ 

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£ 

u u u 



u 

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U 

U 


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O 


o 

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o 

o 


o 

o 


in 


ID 

00 

CO 

00 


00 

• 


• 


• 

• 

• 

• 


• 



CM 


iH 

CO 

rH 

CM 


CM 




0 










G 










rH 







CM 


CM 

0 

rH 



CM 


r- 

in 


in 

> 




LO 


in 




0 







• 

5 


Cn 




• 

— 


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LT> 

in 


0 




CO 

CO 


UO 

rH 



P 




1—1 

rH 


iH 

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— 


0 




— 

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CM 


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cn 


UO 

cn 

rH 

o 


0 




o 

O 

o 

o 

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o 



Pi 



o 

o 

o 

O 

cn 

ID 


'G 

0 



cn 

ID 

cn 

ID 

CO 

r- 


G 

0 



CO 

r- 

CO 

r- 




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P 









to 

g U -H 










P CO 


i 




0 


O 

0 

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rH 

p 




U 


o 

u 

(h 0 • 

G cn 

0 0 

r- 


ID 

0 


0 

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cn -P 

0 Pi CM 

■P Cn 




G 

TS -P 

G -P 

G 

0 P to 

Cn 0 

0 G TS 

0 


0 

G 

G 0 

G 0 

G 

>i 0 

P 0 

0 £ -H 

-P 


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0 

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0 0 

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P £ -P 

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•H 


•H 


to H 

Eh £ 

Eh 

IP 0 

£ u 0 to 

U -P CQ 

CO 


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73 


(continued) 




















TABLE 17. (continued) 


w 

Q 

\ 

C4 

sc 

w 

Q 


[" 

o 



o 



g 

a 

a 


o 

o 


* 

CO 

G 

o 

•H 

P 

fd 

g 

4-> 

c 

Q) 

u 

c 

o 

u 

CD 

P 

fd 



g 

04 04 

SC 04 

w 

p 


co 

O 

• 

o 


G 


04 04 

PQ 04 

Q 


m 



in • 
• o 
o 



o 

UO 

o 

o 


CM 

ID 

00 

• 

o 


CM 

o 

o 


co 


+1 cm 

LD O 

o • 
. o 
o 


i" 

o 


co 

+ I CM 
rH O 


• o 


uo 

+1 «—1 
r- co 
co o 
o • 
• o 


o 


o 


co 

+ I CO 
CO CM 

^ • 
• o 
o 




+1 co 
in • 
• o 
o 


co 

+ |cr\ 
t" o 
in . 

• o 
o 


r" 

o 


in 

+ It" 

CO rH 
CO O 

o • 
• o 
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fd 

p 

4-) 

P 

04 

rH 

>1 

p 

I—I 

< 


c 


g 

04 04 
W 04 

Q 


CM 

CM 

■O’ 

O 

o 



co 

+ I CM 

in o 
cm c 


CM 


+ I CM 

in o 


co 

+1 1 1 
in o 
id o 


CM 

CO 

CM 


o 


o 


o 


o 


• o 


o 


o 


o 


o 


o 


o 



o 

p 

P 

CD 

s 


I" 

I 

CO 


co 

s 

u 

u 


CO 

s 

u 

o 


co 

s 

u 

O 


CD 

P 

•H 

CO 


co 

S 

u 

o 


CO 

s 

u 

o 


cn 

s 

u 

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u 

o\o 


co 

CM 


CM 


CM 


0 

G 

0 

N 


co 


<T> 

CM 


CO 

CM 


o 

CM 

K 


o\° 


Co 

G 

0 

P 

\ 

4-4 

fd 

p 


rH CO 

uo in 


co 


CM 

m 


uo 

o 

o 

<y\ 

co 


00 


00 

o 

o 

CO 

t" 


uo 


uo 

o 

o 

co 

co 


co 

o 


o 

CD 

r- 


CM 


O 

o 

CO 

CO 


uo 

in 


CO 

O 

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CD 

t" 


>1 
P 
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1—I 
fd 
P 

u 

0 

g 


G 

•H 

<D 

G 

rH 

fO 

> 


00 

uo 


t" in 

in cd 


- 

» 

= 

= 

CO 

UO 

rH 

0 

1—1 

uo 

CO 

uo 

CM 

CO 

CO 

CM 

O 

0 

uo 

1—1 

O 

0 

0 

0 

CO 

CD 

00 

CD 

CO 

l" 

CO 

f" 


in 

CM 

o 

co 

co 


o 

v 


4-> 

G 

a) 

g 

•H 

TS 

(D 

in 

Sh 

<D 

> 

•H 

P-1 

Sh 

CD 

P 

cn 

0 

P 

U 

G 

•H 

T5 

G 

G 

0 

P 

P 

CM 0 
A G 


5 h G 

0 15 

44 


G 

0 

> 

•H 

>1 

rH 

G 

0 

0 

fd 

cn 

0 

G 

rH 

> 


0 

p 

rd 

rH 

fd 

P 

p 

fd 

P 

04 


>1 

p 

u 

o 

I 

G 

I 

•H 

a 


0 

p 

•H 

CO 


U0 




CO 


0 

p 

•H 

cn 


0 

P 

•H 

CO 


0 

P 

•H 

CO 


0 

Co 

fd 

H 

0 

> 

< 


CM 

0 

P 

•H 

CO 


0 

P 

•H 

CO 


U 

(X 0 
P H P 
0 cn 0 p 
P 0 > G 
•H P -H 0 
CO u PS S 


0 P4 

Sh o 
U G 
0 Q 

+- + I 


74 


Reagent blanks measured: DEP<0.0005 ppm, DBP<0.001 ppm, DEHP<0.0003 ppm 

* Other alkyl phthalates found in Tenneco Pond (East) DIDP(1.4+0.2x10^(4); 
DTDP(1.9+0.2)xl0 3 (4) 























ANALYSIS OF PHTHALATE ESTERS IN OYSTER TISSUE 


Procedure 


1. Whole oysters were collected from the Chester River and 
pooled according to sampling site. The number ranged 
from 4 to 8 individual oysters per site. 

2. The pooled tissue was homogenized with a Virtis "23" 
homogenizer for one minute at low speed followed by 
two minutes at medium speed. 

3. The homogenized tissue was then disrupted with a 
Branson W185 ultrasonic probe for 10 minutes. 

Light microscopy showed that this caused complete cell 
disruption. 

4. The ultrasonic probe was cleaned by running the device 
twice in distilled water and once in methylene chloride. 
The homogenizer flask and blades were then rinsed with 
water, methanol, and methylene chloride. Blanks of 
water were run between tissue preparation to check for 
contamination. 

5. Twelve to fifteen g of tissue from each site plus two 
controls prepared from commercial oysters were placed 
in 50-ml Erlenmeyer flasks, shell frozen, and placed 
in a vacuum desiccator. 

6. The desiccator was attached to a Virtis lyophylizer and 
the dry tissue was dried in 24 hours. 

7. The dried tissue was removed from each flask, pulverized 
with a mortar and pestle/ and stored at 3°C. 

8. Two hundred mg of dried tissue was placed in 10-ml vials. 
A 10 yg aliquot of anthracene d-10 was added with a 
Corning disposable micro pipette (+0.5 oercent ac¬ 
curacy) . Five ml of methylene chloride was added. 

9. The tissue was extracted using ultrasonic agitation 
(Bransonic 220 water bath) for 5 min. 

10. The vials were centrifuged at 4,000 rpm for 15 min. to 
sediment the remaining tissue residue. The supernatant 
was drawn off. The sample was concentrated by solvent 
evaporation to a volume of approximately one milliliter. 

11. The concentrated extract was injected directly in the 
GC-MS with a 25M capillary column coated with SE-52. 


75 



Samples 




Oyster samples from the Chester River were collected July 2, 
1979 by personnel of the Chesapeake Biological Laboratory. The 
samples were frozen for subsequent delivery. They arrived in 
jars which had cracked presumably during the freezing process. 
The extent of possible contamination, however, was considered 
minimal. 

Oyster sample CBL-1 was collected at Buoy Rock correspond¬ 
ing to sediment site 1. Sample CBL-2 was from Spaniard Point 
where sediment sample 6 was collected. There is no equivalent 
sediment site for the CBL-3 sample collected at Ferry Bar. A 
fourth sample from Love Point contained too few oysters to 
analyze. A homogenate of seven commercial oysters from northern 
Chesapeake Bay was used for reference purposes. Unspiked sam¬ 
ples as well as samples spiked with DEP, DBP and DEHP were pre¬ 
pared from this. 

Results 


Oysters contain approximately 80 percent water and 2 percent 
lipid. Both of these tend to interfere with the analytical de¬ 
termination. The water must first be removed before considering 
extraction of the phthalates with methylene chloride. The lipid 
content is readily soluble in methylene chloride, as are the 
alkyl phthalates. An attempt was made to dry the oyster tissue 
using the same method successfully used for wet sediment. The 
tissue homogenate was spread as thin layer on a watch glass and 
placed in a desiccator containing sodium sulfate. This method 
was discarded because a bacterial bloom tended to form on sam¬ 
ples prepared in this way. However, the controls from commer¬ 
cial oysters were free from this mold-like growth. It is 
suspected that these oysters are thoroughly rinsed and an 
anti-bacterial preservative is added during processing. 

A more rapid drying method was needed. The preferred alter¬ 
native was lyophilization or "freeze-drying." Spiked and un¬ 
spiked controls were first freeze-dried to evaluate the method. 
The spiked samples contained 20 ppm each of DEP, DBP and DEHP. 
The extracts of these samples were injected directly into a 
glass capillary gas chromatograph with an FID detection. 
Numerous, partially resolved peaks resulted but the spiked peaks 
were not evident. GC-MS was used to identify the four major 
sets of peaks. The first large peak was identified by matching 
its mass spectrum to reference spectra as palmitoleic acid and 
closely related compounds that elute at 200°-203°C. The second 
large set of peaks eluting at 215°-217°C was identified oleic 
acid. The third (228°-232°C) and fourth (244°-248°C) groups are 
linoleic acid and linolenic acid, respectively. 


76 




The extract were analyzed for phthalate ester using the 
selected ion monitoring mode. Chromatograms of the spiked and 
unspiked oysters are shown in Figures l 4 and 15. Even though 
percent abundance of m/e 149 is linolenic acid is 3 percent, its 
high concentration in the injected extract makes it clearly 
evident in the m/e 149 selected ion monitoring team. Indeed, 
all four sets of peaks similarly appear in the m/e 149 mass 
chromatogram. While the DEP and DBP are resolved from inter¬ 
fering peaks, DEHP co-elutes with the linolenic acid group. 

The DEHP concentration must be greater than 2 ppm in order to 
make the interfering background insignificant and obtain mean¬ 
ingful quantitative results. 

The recoveries of DEP, DBP and DEHP from spiked (20 ppm) 
oyster tissue takers through the entire drying and extraction 
procedure were 60+6, 96+5, and 100 + 15 percent, respective¬ 
ly. The smaller recovery for DEP is assumed to be due to voli- 
tization during the freeze drying process. 

The measured levels of alkyl phthalates in oyster tissue are 
reported in Table 11 along with values for sediment samples 
taken from the same zone. 

As discussed earlier, the octanol-water partitioning model 
predicts that the hydrophobic phthalates will be concentrated 
in the total lipid portions of the oysters relative to the sur¬ 
rounding water. We did not measure lipid content per se , but 
the oyster and sediment values were compared on the basis of 
the measured total organic carbon content which was measured. 

The oyster contained a 16-fold higher concentration of organic 
carbon than the sediment. Accordingly, the alkyl phthalate 
concentration is predicted by the partition model to be approx¬ 
imately 16-fold higher than that in the sediment. This pre¬ 
diction does not consider any mechanism other than simple 
partitioning. Table 17 shows that the oyster-to-sediment ratio 
of total phthalate ester residue is consistent with this model. 
At the two sites where contiguous oyster and sediment samples 
were obtained, the ratio is 16:1 (Buoy Rock) and 23:1 (Spaniard 
Point). This suggests that the surficial analysis of stratified 
(anoxic, nonbioturbated) taken nearby oyster beds may provide a 
basis for predicting the concentration of phthalates in the 
nearby oysters. It is also apparent that the level of phtha¬ 
lates in the oysters from the Chester River do not signifi¬ 
cantly differ from the reference sample of commercial oysters, 
also taken from the northern Chesapeake Bay. 

Table 18 shows that the measured levels of alkyl phthalates 
in oysters range from 0.45 to 1.5 ppm (wet basis). The U.S. 
Bureau of Sport Fisheries and Wildlife reported recently (Mayer 
et al. 1972) a range of 0.2 to 3.2 ppm of alkyl phthalates in 
channel catfish and walleyes from various parts of North 
America. Also, a survey of 145 catfish farms (Haudet 1970) 


77 





Figure 15 . Oyster tissue extract (spiked with 20 ppm DEP, DBP, and DEHP) 














































co 

■*- 

3 

C 


E 


LU 

1 


79 


Figure 16. Oyster tissue extract. 
































TABLE 18. CONCENTRATION OF PHTHALATES IN OYSTERS AND RELATED SEDIMENTS* 


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81 





















revealed that 95 percent of the fish analyzed contained DEHP 
residues with an average DEHP concentration of 3.15 ppm. 

The alkyl phthalate measurements reported here can be com¬ 
pared, at least indirectly, to available toxicity data. Chem¬ 
ical toxicity results for aquatic organisms are usually 
presented as LC^q values (LC^g is the estimated concentration 
where the toxic substance in the surrounding water will kill 
half the population of exposed organisms within a certain time 
period, usually 96 hours.) Since the present study presents 
values for sediment concentrations, not water, an estimate of 
the water concentration using the octanol/water partitioning 
model was made. The partitioning constant (Kow) of phthalate 
esters is estimated to be 10 3 - 10 4 (TSCA 1978). The average 
sediment values on an organic carbon basis (C oc ) for the mor¬ 
tality zone are 2 ppm, 20 ppm and 2 ppm (yg/g organic acid) for 
DEP, DBP, and DEHP, respectively. The estimated ester concen¬ 
trations in water are, therefore, 0.2 ppb, 2 ppb, and 0.2 ppb 
(yg/L), respectively, for DEP, DBP and DEHP. Mayer and Sanders 
(1973) reported that acute 96 hours LC 50 values for DBP with 
fathead minnor, channel catfish, rainbow trout, scud and cray¬ 
fish fell between 730 and 10,000 ppb (yg/L). The LC^ concen¬ 
trations for DEHP were estimated to be above 10,000 ppb. The 
estimated water concentrations in the Chester River are cur¬ 
rently significantly less than these values. In addition, the 
average alkyl phthalate concentrations in oyster tissue in the 
Chester River range from 3 to 8 ppm. This is 1000-fold lower 
than the acute toxicity of alkyl phthalates for rats (intra- 
peritonel) which ranges from 3-14 g/kg (percent). In comparison, 
the LD 5 o's for organochlorine pesticides in rats are three 
orders of magnitude lower (20 - 300 mg/kg) than alkyl phthalates 
(TSCA 1978). 

Chronic toxic effects of alkyl phthalates on aquatic orga¬ 
nisms appear at much lower levels than acute toxicity. Mayer 
and Sanders (1973) reported that a concentration of only 3 ppb 
of DEHP in the water was sufficient to significantly decrease 
growth and reproduction of the crustacean Daphnia magna . Zebra 
fish and guppy reproduction was also impaired when their food 
was spiked with 50 and 100 ppm of DEHP. Various effects of 
alkyl phthalate have been reported for brine shrimp, goldfish 
and ring doves at concentrations varying from 3 to 10,000 ppb. 

No chronic or acute toxicological data are available on the 
effects of phthalates on oysters. Martin and Roosenburg (1979) 
studied oyster mortality at 10 stations in the Chester River. 

No significant mortality occurred during the four-month period 
of observation. However, during July and August five stations 
furthest upstream had dieoff of the fouling organisms and re¬ 
duced growth rates of the oysters. Ninety-six hour acute 
toxicity studies were performed on golden shiners and crayfish 
in streams receiving effluents from the Campbell's Soup plant. 


82 




the Tenneco, Inc. plant and the Chestertown sewage treatment 
plant. No significant oyster mortality was observed. 

It appears that a comparison of our measurements with the 
known toxic effects of phthalate esters on oysters and other 
aquatic organisms is unable to explain the Chester River oyster 
mortality during 1973-1975. However, there is enough evidence 
to cause concern over long-term effects on aquatic organisms 
especially with expected future increases production of alkyl 
phthalates. 


83 


BIBLIOGRAPHY 


Amundson, S. C. 1978. Determination of di(2-ethylhexyl) 

phthalate, non(2-ethylehexyl) phthalate and phthalic acid by 
high pressure liquid chromatography. 

J. Chromatogr. Sci. 16:170. 

Anonymous. 1976. Outlook for PVC plasticizers in bright. 

Chem. Eng. News (Nov. 22, 1976):12. 

Chesler, S. N., B. H. Gump, H. S. Hertz, W. E. May, S. N. Dyszel, 
and D. P. Enagonio. 1976. Trace hydrocarbon analysis: The 
National Bureau of Standards William Sound/Northeastern Gulf 
of Alasks Baseline Study. NBS Tech. Note 889. 

Gaudet, J. [Ed.] 1970. Report of the 1970 workshop on Fish 

Feed Technology and Nutrition. Bureau of Sport Fisheries 
and Wildlife, Fish and Wildlife Service, U. S. Department of 
Interior. 

Giam, C. S., H. S. Chan, and G. S. Neff. 1975. Sensitive 

method for determination of phthalate ester plasticizers in 
open ocean biota samples. Anal. Chem. 47:2225-2229. 

Giam, C. S., H. S. Chan, T. F. Hammergren, and G. S. Neff. 1976. 
Sensitive method for determination of phthalate ester plas¬ 
ticizers in open ocean biota samples. 

Anal. Chem. 48:78-80. 

Gingras, S. A. 1979. Analysis of alhyl phthalates and other 
organic substances in the environment. M.S. thesis, Univ. 
of Maryland, Department of Chemistry, College Park, MD. 

Han, G. 1972. The delination of an exclusion area around the 
Chestertown outfall on the Chester River and the Back River 
sewage treatment plant outfall. Chesapeake Bay Institute 
(The Johns Hopkins Univ.) Special Rpt. 26, Baltimore, MD. 

Heilman, M. Y. 1978. Analysis of phthalate plasticizers for 

PVC by liquid chromatography. J. Liq. Chromatogr. 1:491-505. 

Kahn, S. U., and M. Schnitzer. The retention of hydrophobic 
organic compounds by humic acid. 

Geochim. Cosmochim. Acta 36:745-754. 


84 


Karickhoff, S. W., D. S. Brown, and T. A. Scott. 1979. Sorp¬ 
tion of hydrophobic pollutants on natural sediments. 

Water Res. 13:241-248. 

Martin, F. D., and W. Roosenburg. 1979. Evaluation of Chester 
River oyster mortality biotoxicity. EPA Rpt. R805976010. 
Chesapeake Biological Laboratory, Solomons, MD., Ref. No. 
79-103 CBL (unpublished). 

Mayer, F. L., Jr., and H. 0. Sanders. 1973. Toxicology of 
phthalic acid esters in aquatic organisms. 

Environ. Health Perspect. 3:153-157. 

Mayer, F. L., Jr., D. L. Stalling, and J. L. Johnson. 1972. 
Phthalate esters as environmental contaminants. 

Nature 238 (Aug. 18, 1972):411-412. 

Meritt, D. W. 1977. Oyster spat set on natural cultch in the 
Maryland portion of the Chesapeake Bay (1939-1975). 

Univ. Md. Ctr. Environ. Estuarine Stud. Spec. Rpt. 7. 

Neely, W. B., D. R. Benson, and G. E. Blau. 197 A . Partition 

coefficient to measure bioconcentration potential of organic 
chemicals in fish. 

Environ. Sci. Technol. 8:1113-1115. 

Palmer, H. D. 1972. Geological investigations, pp. 75-137. 

In: W. D. Clark [Ed.] Chester River study. State of 
Maryland, Dept, of Natural Resources, and Westinghouse Elec. 
Corp., Vol. II. 

Palmer, H. D., J. R. Schubel, and W. B. Cronin. 1975. Estuar¬ 
ine sediraentology, pp. 4/29-4/30. In.: Munson, T. 0., D. K. 
Ela, and C. Rutledge, Jr. Upper Bay survey and final report 
to Maryland Department of Natural Resources, Annapolis, MD 
21401 [Nov. 30, 1975]. 

Pritchard, D. W. 1967. Observations of circulation in coastal 
plain estuaries, pp. 37-44 . In.: G. H. Lauff [Ed.] 

Estuaries. Publ. 83, Amer. Assoc. Advancement Sci., 
Washington, D. C. 

Singmaster, J. A., and D. G. Crosby. 1976. Plasticizers as 
interferences in pollutant analysis. 

Bull. Environ. Contam. Toxicol. 16:291-300. 

U. S. Department of Agriculture. 1973. Composition of foods. 
Agricultural Res. Serv. Handbook No. 8, Washington, D. C. 

U. S. Environmental Protection Agency. 1978. Initial report of 
the TSCA Interagency Testing Committee to the Administrator, 
Environmental Protection Agency, pp. II-i-II-55. 

U. S. EPA 560-10-78/001. 


85 


Watson, J. T. 1976. Introduction to mass spectrometry: 
Biomedical environmental and forensic applications. 

Chpt. 3. Raven Press, N. Y. 

Webster, R. D. J., and J. Nickless. 1976. Problems in the 
environmental analysis of phthalate esters. 

Proc. Anal. Div. Chem. Soc. 13:333-335. 

de Zeeuw, R. A. 1975. Plasticizers as contaminants in high- 
purity solvents:A potential source of interference in bio¬ 
logical analysis. Anal. Biochem. 67:339-341. 


36 


SECTION 6 


MICROBIAL TRANSFORMATION OF TIN 


EXPERIMENTAL PROCEDURES 
Sampling 

Samples were taken in the Chester River at Buoy Rock (lati¬ 
tude 38°59'33"N, longitude 76°12'27"W), which is a productive 
bar; at Spaniard Bar (latitude 30 o 05'57"N, longitude 76°08 1 55"W), 
which has experienced extensive oyster mortality starting in 
1974; at the outfall from the Tenneco plant near Chestertown and 
in the Tenneco holding pond near the pond's outlet (latitude 
39°15'00"N / longitude 76°02'30"W); at the outfall from the 
Chestertown sewage treatment plant at the edge of Morgan Creek 
(latitude 39°12'00"N, longitude 76°04'15"W); and at the Campbell 
factory near Chestertown (latitude 39°14'03"N, longitude 
76"12'21"W). Three sampling excursions were taken in the period 
September 1977 through July 1979. 

For comparison, some samples were taken in Baltimore Harbor 
(latitude 39°13'57"N, longitude 76°30'16"W), which was expected 
to be polluted with heavy metals, and other samples were taken 
near Tilghman Island (latitude 39°40'64"N, longitude 76°23'16"W), 
an area expected to be relatively free of pollution by heavy 
metals. Samples from the two Chester River sites, from Baltimore 
Harbor, and from the site near Tilghman Island were considered 
estuarine samples; samples from other sites were considered fresh¬ 
water samples. 

At each site, water temperature, pH, salinity, and dissolved 
oxygen were determined. Methods are given in Appendix C. 

Water samples were taken with a Kemmerer bottle. Concentra¬ 
ted HC1 (0.5 ml) was added to each 200-ml water sample to keep 
metals in solution. This brought the pH to 1.8 to 1.9. Prior 
to chemical analysis, the pH of each sample was adjusted to 2.0 
with 1 N NaOH. Sediment samples were collected with a Van Veen 
dredge. A plastic corer was used to obtain material which had 
not touched the metal walls of the dredge. Samples were taken 
from the top centimeter of the core, and the remainder of the 
core was sliced into 1-cm bands which were frozen and will be 
maintained frozen for possible future use. 


87 



Samples from the sewage treatment plant, from the Campbell 
factory, and from the Tenneco plant were iced in the field and 
stored in ice until they were used in the laboratory. Samples 
taken on board ship (Buoy Rock, Spaniard Bar, Baltimore Harbor, 
Tilghman Island) were used for microbiological analysis within 
15 min of sampling; the remainder of each sample was stored on 
ice until it was used in the laboratory. 

Microbiological Samples 

Water samples were used to prepare appropriate dilutions for 
plating. For sediment samples, 1.0 g (wet weight) was suspended 
in 9.0 ml of sterile estuarine salts; further dilutions were pre¬ 
pared from this suspension. 

Total viable counts of aerobic, heterotrophic bacteria were 
made using the spread plating technique, plating on Nelson's 
medium (Nelson et al. 1973). Nelson's medium contains casamino 
acids; 5.0 g; yeast extract, 1.0 g; glucose, 2.0 g; agar, 15.0 g; 
and salt solution, 1 liter. For sediment samples from estuarine 
sites, the estuarine salts solution contained NaCl, 10.0 g; 
MgCl2*6H20, 2.3 g; KC1, 0.3 g; and distilled water, 1 liter. 

For samples from freshwater sites, the salts solution was used 
at one-tenth strength. 

For viable counts of tin-resistant organisms, appropriate 
dilutions were spread on the surface of Nelson's medium prepared 
as above, supplemented with SnCl^ to yield 75 ppm tin suspended 
in the medium. Extensive preliminary testing (Table 19 ) indi¬ 
cated that this concentration of tin was appropriate to select 
for tin-resistant organisms. Addition of SnCl^ to the medium 
resulted in a fine precipitate of SnC >2 which was uniformly sus¬ 
pended in the medium by agitation. In one series of experiments 
the organisms resistant to organotin were estimated by plating 
on Nelson's medium containing 15 ppm tin as (CH 3 ) 2 SnCl 2 . 

All platings were prepared in triplicate. Plates were incu¬ 
bated at 25 + 2 C for 3 days prior to counting time. 

Two systems were employed to determine if the microbial flora 
in samples could transform tin to volatile organotin compounds: 

i) Bioflasks . 250-ml flasks containing 20 ml of Nelson's 

medium supplemented with 75 ppm tin as SnCl^, were inoculated 
with 1.0 ml of sediment suspended in estuarine salts. Each 
flask contained a 10-ml beaker embedded in the agar.^ The beaker 
held 5.0 ml of a solution of 8 percent (wt/vol) citric acid in 
10 percent HC1. The flask was sealed with a rubber stopper. 

After 14 days incubation at 27 + 2 C, material in the beaker was 
examined for tin. Thus, if organisms growing on the medium pro¬ 
duced volatile tin compounds, they would be detected in the acid 
solution. Sterile controls to check for nonbiological production 


RR 





TABLE 19. DETERMINATION OF A CONCENTRATION OF TIN WHICH 
WOULD SELECT FOR TIN-RESISTANT MICROORGANISMS 


Type of 
Sample 

Tin added 
(ppmSn, as 
SnCl 4 ) 

Viable count 

(mean 

+ standard 

Water 

0 

8.7 

X 

10 2 

+ 

1.8 

X 

10 2 a 


50 

5.4 

X 

10 2 

+ 

6.8 

X 

ioi b 

Sediment 

0 

8.1 

X 

10 5 

+ 

2.6 

X 

10 4 a 


50 

9.4 

X 

10 4 

+ 

8.5 

X 

10 3 b 


100 

4.0 

X 

10 4 

+ 

3.9 

X 

10 3 c 


150 

2.6 

X 

10 4 

+ 

7.1 

X 

10 3 c 


200 

1.7 

X 

10 4 

+ 

3.4 

X 

10 3 c 


* 

Means with the same superscript are not significantly different 
at the 5 percent level as determined by a one-way analysis of 
variance (ANOVA). 


of volatile tin compounds were included in each experiment. 

Each experiment also included positive controls in which the 
medium contained 75 ppm tin as dimethyltin chloride. The use of 
a solid medium containing a suspension of tin renders this method 
qualitative, not quantitative. 

ii) Hungate tubes . Each tube (Hungate 1969) , containing 
5.0 ml of liquid Nelson's medium (Nelson's medium minus agar), 
was inoculated with 1.0 ml of sediment suspended in estuarine 
salts. Sterile controls and positive controls were included in 
each experiment. Tubes were incubated for 16 days at 27 + 2 C 
on a rotary shaker operating at 80 rpm; each tube was then sam¬ 
pled for the presence of organotin compounds. 

For each water or sediment sample, one set of bioflasks and 
Hungate tubes was inoculated from a sample which had received 
500 pg of sodium azide per ml. This additional set of controls 
was used as a demonstration that effects observed in cultures 
were due to biological activity. 

Analyses of Tin 

Inorganic tin— 

For sediment samples, 1.0 g (wet weight) was transferred to 
an acid-cleaned, screw-capped tube. Then, 2.0 ml of a solution 
containing 50 percent (vol/vol) HC1 and 50 percent (vol/vol) 
concentrated HNO 3 were added. The tube was shaken vigorously 
for 1 hr. It was then centrifuged and 1.0 ml of the supernatant 


89 










fluid was removed for analysis. The method of additions (Perkin- 
Elmer 1977) was used to minimize matrix interference. 

Water samples (100 ml, not filtered) were adjusted to pH 2. 

A 15.0-ml quantity was then extracted with 10 ml of 15 percent 
(wt/vol) ammonium pyrrolidine dithiocarbamic acid (APDC) in dis¬ 
tilled water. The APDC solution was prepared by dissolving APDC 
in distilled deionized water which had been adjusted to pH 7 
with NaOH. The APDC solution was then extracted three times with 
CCI 4 to remove impurities. The water sample, containing APDC, 
was stirred for 10 min. It was then extracted with a minimum 
volume of methyl isobutyl ketone (MIBK). The mixture was agita¬ 
ted for 1 min and then allowed to separate for 10 min. The MIBK 
phase was analyzed for tin. 

Organic tin-- 

The organic acid solution from bioflasks was analyzed di¬ 
rectly for tin content. 

Medium from Hungate tubes was extracted in such a way that 
inorganic tin compounds remained behind while organotin com¬ 
pounds were extracted. Extensive preliminary experiments showed 
that this extraction was effective. Medium was centrifuged at 
3000 x g for 10 min to remove cells and sediment. The super¬ 
natant medium was then extracted with 2.0 ml of dichloromethane: 
chloroform (9:1, vol/vol). The extraction was conducted over a 
period of 1 hr, with periodic agitation. The mixture was then 
centrifuged at 1,000 x g for 10 min to sediment remaining in¬ 
organic tin precipitate. The lower, organic phase was removed 
and evaporated by dryness under a stream of nitrogen. The re¬ 
sidue was dissolved in MIBK and analyzed for tin. Evaporation of 
the organic phase under nitrogen may have removed some organotin 
material, since organotins are volatile. Thus, results from Hun¬ 
gate tubes are qualitative, not quantitative. 

Atomic absorption spectrophotometry— 

Tin was analyzed with the aid of an HC-2200 graphite furnace 
with ramp accessory on a Perkin-Elmer model 503 AA unit using D 2 
background correction (to minimize interference from "smoke" and 
nonatomic absorption) and an Sn electrodeless discharge lamp 
(EDL). Argon was the sheath gas. 

Mineral acid solutions from sediment samples and organic 
acid solutions from bioflasks were analyzed with the EDL set at 
286.3 nm. Samples were dried at 105 C for 40 sec (10 sec ramp), 
charred at 800 C for 40 sec (10 sec ramp) , followed by atomiza 
tion using maximum power at 2700 C for 8 sec with a 3-sec stop 

of gas. 

MIBK from water samples and from biotubes was analyzed with 
the EDL set at 224.5 nm. Samples were dried at HOC for 40 sec 


90 


(10 sec ramp), charred at 650 C for 40 sec (no ramp), followed by 
atomization at 2500 C for 8 sec with a 3-sec stop of gas. 

RESULTS AND DISCUSSION 

Partially Developed Methods 


One of the objectives of this work was to apply quantitative 
methods developed for other metals and organometallic compounds 
to tin and organotin compounds. The approach was to separate 
organotin compounds from one another via gas liquid chromato¬ 
graphy (GLC) and to allow the effluent from the GLC to pass 
through a heated transfer tube to an atomic absorption spectro¬ 
photometer (AA) which would detect tin. Thus, those compounds 
which contained tin would be detected by the AA. The method is 
in use in several laboratories for other metals (Brinckman et 
al. 1976, Parris et al. 1977, Trachman et al. 1977) and contact 
was maintained with workers at the National Bureau of Standards 
throughout the project. Due to time pressures, further develop¬ 
ment of the method was stopped before it was ready for use on 
this project. Success was achieved in separating mono-, di-, 
tri-, and tetra-methyltin via GLC, although the system is not 
yet sensitive enough for direct application to environmental 
samples. The A?, unit is close to being used as a detector for 
organotin compounds eluted from the GLC. Not the least of the 
difficulties was an 11-month delay in receiving the heated 
transfer line from the sole manufacturer. Work on this aspect 
of the project is continuing and support of this contract will 
be acknowledged in all publications. 

Physical and Chemical Data on Samples 


Physical and chemical data for samples taken on cruises and 
excursions in April 1978 and July 1979 are summarized in 
Tables 20 and 21. Temperatures and salinities were as expected 
for these sites in spring and summer seasons, respectively. pH 
values, which were taken only for the summer 1979 samples, were 
significantly higher at the freshwater sites than at the estu¬ 
arine sites; they were highest at the two Tenneco sites. Values 
for dissolved oxygen were higher in April 1978 than in July 1979, 
as expected for spring and summer seasons, respectively. It is 
noteworthy that in summer 1979, dissolved oxygen was dangerously 
low near the bottom at the Buoy Rock site, which did not suffer 
extensive oyster mortality; while dissolved oxygen was only 
slightly higher at Spaniard Bar. Both oyster bars gave lower 
summer readings for dissolved oxygen than were found in Balti¬ 
more Harbor. A very high reading was obtained at the Tenneco 
pond in July 1979. A comparison of physiochemical data for the 
healthy bar. Buoy Rock, and the data for Spaniard Bar, show only 
dangerously low dissolved oxygen near the bottom as a potential¬ 
ly lethal condition. None of the physiochemical data from the 
freshwater sites gives a direct clue to the extensive kill of 


91 




TABLE 20. 

PHYSICAL DATA 

FOR APRIL 1978 CRUISE/EXCURSION 


Station/Site 

Depth* 

(meters) 

Temperature Salinity 

<° C) (Voo) 

Dissolved 
Oxygen 
(parts per 
thousand) 


Buoy Rock 

1 

10.8 

4.4 

10.6 


6 

11.7 

5.2 

10.7 


12 

7.7 

8.1 

10.2 

Spaniard Bar 

1 

12.3 

4.6 

10.3 


5 

12.0 

5.5 

10.3 


10 

12.1 

5.4 

10.2 

Tenneco - effluent 

— 

20 

3 

9.1 

Tenneco - pond 

0.2 

17 

2 

10.1 

Campbell plant 

0.2 

18 

1 

7.0 

Chestertown sewage 





treatment plant 

0.2 

19 

1 

9.4 


TABLE 21. PHYSICAL DATA FOR JULY 1979 CRUISE/EXCURSION 

Station/Site Depth* Temperature pH Salinity Dissolved 

(meters) (° C) (°/oo) Oxygen 


(parts per 
thousand) 


Buoy Rock 

1 

22.6 


6.5 

6.2 


6 

22.5 


6 .6 

6.0 


12 

22.0 

6.6 

6.8 

4.5 

Spaniard Bar 

1 

24.0 


5.6 

7.4 

5 

23.9 


5.6 

7.4 


10 

23.9 

6.1 

5.6 

7.3 

Baltimore Harbor 

1 

23.2 


3.5 

7.9 


2.5 

22.8 


3.5 

8.0 


5 

22.7 

5.2 

3.8 

7.6 

Tilghman Island 

1 

22.5 


8.7 

9.4 

5 

22.6 


8 .8 

9 .6 


10 

22.6 

5.5 

9.1 

9.3 

Tenneco - effluent 

— 

30.0 

9.6 

2.0 

8.4 

Tenneco - pond 

0.2 

24.9 

8.4 

2.0 

>20 

Campbell plant 

0.2 

21.0 

7.2 

2.0 

9.5 

Chestertown sewage 


24.0 


2.0 

A 

treatment plant 

0.2 

7 .6 

9 . 4 


The deepest reading at each point was taken at approximately 
1 m from the bottom. 


92 











benthic fauna in the Chester River. It is interesting that dis¬ 
solved oxygen values for Baltimore Harbor are lower than for the 
site at Tilghman Island and only slightly higher than the values 
at Spaniard Bar. 

Data for Microbial Populations 


Data for enumeration of bacteria are shown in Tables 22 and 
23. Counts in sediment are higher than counts in water, as ex¬ 
pected. In general, estuarine sites showed higher bacterial 
counts in summer than in spring; this is expected as a function 
of temperature and of increased production of organic materials 
in the estuary in the summer. 


TABLE 22. VIABLE COUNTS OF BACTERIA FROM APRIL 1978 CRUISE 


Station/Site 

Type of 
Sample 

Total 

Viable 

Sample 

Resistant to 
Inorganic-Sn 

% 

Resistant 

Buoy Rock 

Water 

1 .2xl0 2 

1 .6xl0 2 

133 


Sediment 

2.OxlO 5 

1.3xl0 4 

6 

Spaniard Bar 

Water 

<1.2xl0 2 

<1.2xl0 2 

_* 


Sediment 

2 .4xl0 5 

1 .2xl0 5 

50 

Tenneco - 
effluent 

Water 

2 .8xl0 4 

2.OxlO 4 

10 

Tenneco - 

Water 

l.lxlO 4 

2 .2xl0 3 

20 

pond 

Sediment 

2.4xl0 6 

3.7xl0 5 

16 

Campbell 

Water 

6 .4xl0 3 

3.2xl0 3 

50 

plant 

Sediment 

4.lxlO 7 

2.lxlO 7 

51 

Chestertown 

Water 

1 .4xl0 2 

<1.2xl0 2 

_ * 

sewage 

treatment 

plant 

Sediment 

5.8xl0 5 

3.2xl0 5 

55 


* 

Data not accurate enough to yield a useful percentage. 


93 








TABLE 23. VIABLE COUNTS OF BACTERIA FROM JULY 1979 CRUISE 


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94 












In most cases, a significant fraction of the microbial popu¬ 
lation was resistant to inorganic tin and is, therefore, poten¬ 
tially capable of metabolizing tin to more toxic compounds. A 
much smaller fraction of the population was resistant to the 
organotin compound, dimethyltin chloride, attesting to the anti¬ 
bacterial properties of organic tin compounds. The data are con¬ 
sistent with the hypothesis that aquatic microorganisms can 
protect themselves against toxic tin compounds by transforming 
tin to organotins which, although toxic in themselves, are vola¬ 
tile and leave the immediate vicinity of the cell which formed 
them. 

Sediment from Spaniard Bar did not contain higher numbers of 
tin-resistant organisms than sediments from Buoy Rock, although 
a higher percentage of the population was resistant to tin at 
Spaniard Bar than at Buoy Rock. Sediment samples taken at the 
Chestertown sewage treatment plant contained a high percentage 
of organisms resistant to inorganic tin. There is little indi¬ 
cation of a higher level of tin-resistant microflora at Spaniard 
Bar than at Buoy Rock. The data available do not suggest that 
the freshwater sites contain higher numbers of tin-resistant or¬ 
ganisms than the estuarine sites. 

Data for production of volatile tin compounds are summarized 
in Table .24. The method used was effective in detecting organic 
tin compounds produced in bioflasks, as indicated by values ob¬ 
served from sterile medium which contained dimethyltin chloride. 
Variation among replicates indicates that the method is qualita¬ 
tive. Flasks which received inoculum containing the metabolic 
poison sodium azide yielded no volatile tin, indicating that 
volatile tin detected was the result of biological activity. 
Results from bioflasks and Hungate tubes demonstrate that each 
site contains microorganisms capable of converting inorganic tin 
to volatile organotin(s). The species of organotin produced were 
not identified. 

Tin in Water and Sediments 


The lower limit of sensitivity for tin analysis was 2 ppb. 

A recovery value of 77.8 percent was obtained when sediment from 
Tilghman Island was spiked with SnCl/j. When salt water or 
Nelson's liquid medium was spiked with SnCl^, recoveries of 
94-96 percent were obtained. 

Sediment samples contained more tin than water samples 
(Table 25), as expected. Sediment from Baltimore Harbor, known 
as a polluted site, contained over 200 ppm tin (>0.02 percent on 
a wet weight basis). In contrast, sediments from the Tilghman 
Island site contained less that 1 ppm (<0.0001 percent on a wet 
weight basis). All sediments associated with the Chester River, 
including sediments from the three freshwater sites, yielded more 
tin than sediments from the Tilghman Island site (Table 25). 


95 



TABLE 24 . PRODUCTION OF VOLATILE TIN IN MEDIA INOCULATED WITH 



_ Result from _ 

Bioflask Hungate tube 

Replicate yg sn in 

No « organic 

acid 

solution* 


None, sterile control 

1 

148 

++++t 

with medium contain- 

2 

45 


ing dimethyltin 
chloride 




Buoy Rock 

1 

21 

+ 


2 

0 


Spaniard Bar 

1 

262 

++ 


2 

57 


Baltimore Harbor 

1 

344 

+ 


2 

145 


Tilghman Island 

1 

658 

+ 


2 

0 


Tenneco pond 

1 

361 

++ 


2 

314 


Campbell plant 

1 

218 

+ 


2 

144 


Chestertown sewage 

1 

176 

++ 

treatment plant 

2 

201 



Source of 
inoculum 


* Corrected for values obtained from sterile flasks containing 
inorganic tin. 

t The notation used represents peak height from the atomic 
absorption unit: 0.0; +, 0.1 to 0.2; ++, 0.3 to 0.6; 

+++, 0.6 to 1.0; ++++, 1.0 and above. 


96 







TABLE 25 . TIN IN WATER AND SEDIMENT SAMPLES _ 

ug/ml or pg/g for samples taken in 


Station/Site 

Type of 
Sample 

April 1978 

July 1979 


Mean 

Standard 

Mean 

Standard 




error 


error 

Buoy Rock 

Water 

<0.002 

— 

<0.002 

— 


Sediment 

1.574 

0.180 

7.882t 

0.088 

Spaniard Bar 

Water 

0.240 

0.007 

<0.002 

— 


Sediment 

9.686 

0.420 

3.061| 

0.187 

Baltimore 

Water 



<0.002 

— 

Harbor 

Sediment 



239.633* 

14.073 

Tilghman 

Water 



<0.002 

— 

Island 

Sediment 



0.8615 

0.052 

Tenneco - 

Water 

0.034 

0.002 

0.052 

0.002 

effluent 






Tenneco - 

Water 

0.103 

0.005 

0.023 

0.001 

pond 

Sediment 

0.043 

0.081 

3.523;j; 

0.046 

Campbell 

Water 

0.032 

0.001 

<0.002 

— 

plant 

Sediment 

1.583 

0.131 

3.876;j; 

0.028 

Chestertown 

Water 

0.002 

— 

0.152 

0.028 

sewage 

treatment 

plant 

Sediment 

2.857 

0.060 

5.178t, 

t 0.028 


*, + , t, §. Sediment samples values with identical superscripts 
were nbt significantly different at the 5 percent confidence 
level. 


97 









Smith and Burton (1972) reported values of <0.05 to 16 ppm in 
various marine sediments. Surface sediments from Narragansett 
Bay contain 20 ppm tin (Hodge, Seidel, and Goldberg 1979). Furr 
et al. (1976) reported values as high as 492 ppm tin in sewage 

sludge. Thus, values obtained for sediments in the present study 
are in the same range as reported by other workers. Sediments in 
the Chester River, in the Tenneco holding plant, near the Camp¬ 
bell plant, and in the Chestertown sewage treatment plant all 
contain significant quantities of tin (Table 25) . Based on the 
two samples taken, Spaniard Bar, which suffered an oysterkill, 
did not yield significantly more tin than Buoy Rock, which did 
not suffer such a kill (Table-25). The null hypothesis that 
mean tin concentration (Spaniard Point) did not differ from the 
mean tin concentration (Buoy Rock) was supported. Thus, it is 
not possible to attribute the oysterkill at Spaniard Bar solely 
to pollution by tin, although interaction of tin with other fac¬ 
tors cannot be excluded. It is also possible that different tin 
species are present at the two sites, e.g. inorganic tin vs. one 
or more organotin species. 

Smith and Burton (1972) reported inorganic tin concentrations 
of 0.02 to 0.04 yg/kg (0.002 to 0.04 parts per trillion) in 
estuarine and continental shelf waters. Hodge et al. (1979) re¬ 
ported 2 to 38 ng inorganic tin/liter (0.002 to 0.038 yg/kg, 0.002 
to 0.038 parts per trillion) in water from San Francisco Bay and 
San Diego Bay, and 84 to 490 yg/liter (0.084 to 0.490 yg/kg, 

0.084 to 0.490 parts per trillion) in water from Lake Michigan. 

In Lake Michigan water, the concentrations of organic tin com¬ 
pounds were two to eight times greater than the concentrations 
of inorganic tin. The methods used in these two studies were 
several orders of magnitude more sensitive than the method 
employed in the present work. Some of the values obtained for 
water samples in the present work (Table 25) are significantly 
higher than values reported by Smith and Burton (1972) and by 
Hodge et al. (1979). Variations from one sample time to another 
(e.g., values for water from Spaniard Bar and water from the Ten¬ 
neco pond) can be expected due to variations in input and in flow 
rate, which can also account for the low value in the water sam¬ 
ple from Baltimore Harbor. Our data suggest that water in the 
Chester River and water entering the Chester River from the Ten¬ 
neco plant, from the Campbell plant, and from the Chestertown 
sewage treatment plant sometimes contains significant quantities 
of tin. The chemical species of tin were not identified in the 
present study. 

The data obtained in the present study yielded reasonable 
recovery from spiked samples. The method of additions was used 
successfully to alleviate matrix interference. Thus, the values 
obtained for tin are regarded with confidence, even though they 
are high. 


98 



Statistical Analyses 


Data for bacterial enumerations, and data for tin levels were 
analyzed using a one-way analysis of variance (ANOVA). Sample 
means were compared using the studentized (SNK) multiple range 
test. 

Correlation coefficiencts (r) were analyzed for significance 
of association between tin concentration and counts of tin-re¬ 
sistant microorganisms; and for significance of association 
between tin concentration and percent of the microbial popula¬ 
tion resistant to tin. All associations were tested at the 
5 percent level of significance. No significant positive or 
negative correlations were noted. Thus, the data available do 
not suggest that tin (organic and/or inorganic) selects for a 
population of tin-resistant microorganisms. 

General Discussion 


The data indicate that surface sediments from oyster bars in 
the Chester River contain significantly more tin than is con¬ 
tained in sediments from a site near Tilghman Island, but they 
contain significantly less tin than sediment from Baltimore 
Harbor. The three freshwater sites examined--the Tenneco plant, 
the Campbell factory and the Chestertown sewage treatment plant-- 
all contribute tin to the Chester River ecosystem at an undeter¬ 
mined rate. There may be other contributors. It is clear that 
microorganisms capable of converting inorganic tin to more toxic 
organotin compounds are ubiquitous in the Chesapeake ecosystem 
(Table 24) . 

Zuckerman et al. (1978) reviewed the known toxic effects of 
organotin compounds on a variety of living organisms. Relative¬ 
ly little is known of effects on estuarine organisms, but the 
following toxic levels have been reported for organisms relevant 
to the present study: guppies, less than 1 ppm of bis(tributyl- 
tin) oxide and 0.1 ppm triphenltin hydroxide; molluscs, 1.0 ppm 
of several trialkyltin and 0.05 - 0.10 bis(tributyltin)oxide; 
algae, barnacles, shrimp and tubeworms, 0.1 to 1.0 ppm of tri¬ 
butyl and triphenyltin compounds. Very little is known of the 
levels of organotin compounds in estuarine systems. But sedi¬ 
ments in the Chester River ecosystem contain between 3 and 
8 ppm of tin (Table 25). Data from the present study do not in¬ 
dicate the chemical speciation of the tin compound(s) detected. 
Although the chemical species of tin compound (s) was not estab- 
lised, if even one-third of the tin is present as organotin 
compounds, the estuarine biota could be at risk, particularly 
if stress from toxic tin(s) is coupled with other stresses such 
as low dissolved oxygen. 


99 




Comparisons of tin in surface sediments from Spaniard Bar 
with data from Buoy Rock and with data from other marine sedi¬ 
ments (Smith and Burton 1972, Hodge et al. 1979) indicate clearly 
that tin is not the sole source of the oysterkill at Spaniard 
Bar. However, if oysters and other benthic organisms were stres¬ 
sed by other factors, e.g. low dissolved oxygen, tin could con¬ 
tribute additional stress leading to death. 

Little is known about bioaccumulation of tin by benthic 
invertebrates. Until recently, analytical methods were not 
sensitive enough to permit one to deal with tin in the same 
quantitative manner as with other inorganic pollutants such as 
copper, cadmium, mercury, and zinc. These include aspects of 
levels of tin and organotins in the environment, accumulation of 
tin compounds in the food chain, chemical and biological forma¬ 
tion and transformation of inorganic tins and organotins, trans¬ 
port of tin compounds in soils and in aquatic ecosystems, and 
toxicity of tin compounds to the biota. The appropriate ques¬ 
tions should be asked regarding tin. Moreover, it is possible 
that the gut flora of oysters and other invertebrates can con¬ 
vert inorganic tin to more toxic organotins; these questions 
should be addressed. 


100 


BIBLIOGRAPHY 


Brinckman, F. E., W. P. Iverson, and W. Blair. 1976. Approaches 
to the study of microbial transformations of metals, pp. 919- 
936. In : J. M. Sharpley and A. M. Kaplan (Eds.) Proceedings 
of the Third International Biodegradation Symposium. Applied 
Science Publishers, London. 

Daum, R. J. 1965. Agricultural and biocidal applications of 
organometallics. Ann. N. Y. Acad. Sci. 125:129. 

Deschiens, R., and H. Floch. 1962. Comparative study of the 
molluscidial action of 5, 2'-dichloro-4'-nitro-salicyl- 
anilide and salts of triphenyl tin. Bull. Soc. Pathol. 

Exot. 55:816. 

Furr, A. K., A. W. Lawrence, S. S. C. Teny, M. C. Grandolfe, 

R. A. Hofstader, C. A. Bache, W. H. Gutenman, and D. J. Lisk. 
1976. Multielement and chlorinated hydrocarbon analysis of 
municipal sewage sludges of American cities. Environ. Sci. 
Technol. 10:683-687. 

Hodge, V. F., S. L. Seidel, and E. D. Goldberg. 1979. Deter¬ 
mination of tin (IV) and organotin compounds in natural 
waters, coastal sediments and macro algae by atomic spectro¬ 
metry. Anal. Chem. 51(8):1256-1259. 

Holden, A. W. 1972. The effects of pesticides on life in fresh 
waters. Proc. R. Soc. Lond. 180:383. 

Huey, C., F. E. Brinckman, S. Grim, and W. P. Iverson. 1974. 

The role of tin in bacterial methylation of mercury. Proc. 
Int. Conf. on Transport of Persistent Chemicals in Aquatic 
Ecosystems II, pp. 73-78. Natl. Res. Council, Ottawa, 

Canada. 

Hungate, R. E. 1969. A roll tube method for cultivation of 
strict anaerobes, pp. 117-132. In: J. R. Norris and D. W. 
Ribbons (Eds.) Methods in Microbiology, Vol. 3B. Academic 
Press, London. 

Kimmel, E. C., R. H. Fish, and J. E. Casida. 1977. Bioorganotin 
chemistry, metabolism of organotin compounds in microsomal 
monooxygenase systems and in mammals. 

J. Agr. Food Chem. 25(1):l-9. 


101 


Lloyd, R. V., and M. T. Rogers. 1973. Electron spin resonance 
study of some silicon-, germanium-, and tin-centered radi¬ 
cals. J. Am. Chem. Soc. 95(8):2459-2464. 

♦ 

Luijten, J. G. A. 1972. Applications and biological effects of 
organotin compounds, pp. 931-932. In: A. K. Sawer (Ed.) 
Organotin Compounds. Marcel Dekker, New York. 

Nelson, Jr., J. D., W. Blair, F. E. Brinckman, R. R. Colwell, and 
W. P. Iverson. 1973. Biodegradation of phenylmercuric ace¬ 
tate by mercury resistant bacteria. 

Appl. Microbiol. 26:321-326. 

Parris, G. E., W. R. Blair, and F. E. Brinckman. 1977. Chemical 
and physical considerations in the use of atomic absorption 
coupled with a gas chromatograph for determination of trace 
organometallic gases. Anal. Chem. 49:378-384. 

Perkin-Elmer. 1977. Analytical methods for atomic absorption 

spectoscopy using the HGA graphite furnace. 

Perkin-Elmer Corp., Norwalk, CT. 

Ridley, W. P., L. J. Dizikes, and J. M. Wood. 1977. Biomethyla- 
tion of toxic elements in the environment. 

Science 197 (4301) :329-332. 

Smith, J. D., and J. D. Burton. 1972. The occurrence and dis¬ 
tribution of tin with particular reference to marine environ¬ 
ments. Geochim. Cosmochim. Acta 36:621-629. 

Subramanian, R. V. 1978. Recent advances in organotin polymers. 
Poly.-Plast. Technol. Eng. 11 (1):81-116. 

Thayer, J. S. 1974. Organometallic compounds and living 
organisms. J. Organometal. Chem. 76:265-295. 

Trachman, H. L., A. J. Tyberg, and P. D. Branigan. 1977. 

Atomic absorption spectrometric determination of sub-part- 
per million quantities of tin in extracts and biological 
materials with a graphite furnace. Anal. Chem. 49:1090-1093. 

Van der Kerk, G. J. M. 1976. Organotin chemistry:past, present, 
and future, p. 2. In: J. J. Zuckerman (Ed.) Organotin Com¬ 
pounds: New Chemistry and Applications. American Chemical 
Society, Washington, D. C. 

Zuckerman, J. J., R. P. Reisdorf, H. V. Ellis III, and R. R. 

Wilkinson. 1978. Organotins in biology and the environment, 
pp. 388-424. In: F. E. Brinckman and J. M. Bellama (Eds.) 
Organometals and organometalloids, occurence and fate in the 
environment. American Chemical Society, Washington, D. C. 


102 


APPENDIX A 


CHESTER RIVER OYSTER MORTALITY 

A review of recent Fisheries Administration catch records 
and oyster bar surveys of the upper Chester River indicates a 
significant decline in the populations of both oysters and 
associated organisms. 

A major oyster mortality occurred during the spring and 
summer of 1973 in Langford Creek and the Chester River above 
Oldfield oyster bar. The 1974 fall survey showed the mor¬ 
tality was continuing as Oldfield had died-off and some mor¬ 
tality had occurred on Piney Point bar. No live oysters were 
found above Hells Delight bar during the 1975 oyster survey. 

These mortalities caused oyster landings from the area above 
Piney Point bar to decline from 50,000 bushels in 1972-1973 
to 650 bushels for the 1975-1976 season. 

The following is the text of a 8 June 1978 letter sent to 
us by George E. Krantz of the Horn Point Environmental Labora¬ 
tories : 

"Please excuse the long delay in sending you some data on 
the Chester River mortality. Unfortunately my files were loaned 
to other investigators who removed many of the original documents 
that I described to you during our phone conversation last month. 
I think I have found copies of most of the information but I was 
unable to find a complete briefing document for you. Perhaps 
some of the data, especially from the Dept, of Natural Resources 
will be helpful in your study. At a later date we may be able 
to more thoroughly discuss the observations that may have ex¬ 
isted in this historical phenomenon." 

Data on pages 103 thru 110 were compiled by Roy Scott, DNR, 
from field data sheets reflecting Fall oyster bar survey results. 
Time of survey of specific bars varied from October thru March of 

a given year. 


103 


1970. Continuation of ltr 

. from G. 

E. Krantz of June 8, 1978. 


Bar 

Markets* 

Smalls* 

Spat* 

Boxes* 

Ferry 

216/75 

90/8 

2 

4 

Ferry 

174/65 

236/30 

2 

8 

Side Shoal 

42/20 

762/80 

2 

2 

Buoy Rock 

198/70 

74/15 

28 

2 

Buoy Rock 

188/60 

43/15 

4 

0 

Buoy Rock 

110/55 

40/20 

0 

6 

Blunt 

50/9 

0 

2 

58 

Blunt 

180/65 

244/30 

0 

0 

Blunt 

266/90 

12/4 

4 

0 

Hail Point 

114/50 

208/30 

6 

2 

Hail Point 

170/70 

26/5 

0 

0 

Hail Point 

182/85 

12/4 

0 

2 

Hail Point 

98/60 

12/2 

2 

2 

Carpenters Island 

124/40 

530/55 

4 

4 

Carpenters Island 

236/75 

28/3 

4 

4 

Durdin 

98/30 

406/45 

4 

10 

Durdin 

124/40 

152/50 

8 

10 

Durdin 

36/40 

22/1 

2 

2 

Horserace 

182/95 

6 

2 

0 

Horserace 

110/45 

414/40 

4 

0 

Piney Point 

202/70 

38/10 

0 

4 

Bay Bush Point 

160/70 

56/10 

6 

12 

Hells Delight 

158/60 

4 

0 

0 

1 

Hells Delight 

282/90 

4 

0 

4 

(continued) 

104 





1970. 

Continuation of ltr. 

from G. E. Krantz of June 8, 1978. 


Bar 

Markets* 

Smalls* Spat* Boxes* 

3$ 


Bluff Point 

212/80 

28/2 

0 

6 

Middlegrounds 

230/85 

16/2 

0 

10 

Oldfield 

4 

686/85 

0 

16 

Oldfield 

216/75 

16/3 

0 

8 

Oldfield 

156/50 

22 

6 

0 

Willow Bottom 

116/55 

10/2 

0 

10 

Hudson 

186/50 

30/6 

4 

8 

Hudson 

174/30 

494/60 

0 

8 

Boathouse 

78/25 

234/35 

0 

6 

Drum Point 

55/35 

20/3 

0 

0 

Davis Creek 

142/60 

30/3 

0 

4 

Ebb Point 

100/45 

306/38 

4 

2 

Ebb Point 

132/40 

117/12 

2 

2 

Spaniard Point 

148/30 

730/60 

0 

4 

Spaniard Point 

4 

908/100 

0 

0 

Cliff 

72/30 

400/60 

0 

6 

Emory Hollow 

20/15 

2 

0 

2 

Mummys Cove 

116/65 

4 

2 

4 

Shippen Creek 

132/70 

0 

0 

4 

Shippen Creek 

170/50 

18/3 

0 

16 


* 

Number/Percentage of 1 oyster bushel. 


105 







1973• Continuation of ltr. from G. E. Krantz of June 8, 1978. 


Bar 

Markets* 

Smalls* 

Spat* 

Boxes* 

Associated 
Organisms 
Present + 

Buoy Rock 

152/70 

80/10 

0 

0 

0 

Hail Point 

162/90 

22/4 

2 

2 

X 

North of Hail Pt. 

108/50 

36/7 

0 

4 

X 

Carpenters Island 

114/50 

16/2 

0 

4 

X 

Durdin 

188/65 

80/7 

0 

8 

X 

Horserace 

118/50 

58/10 

0 

8 

0 

Piney Point 

54/30 

0 

0 

12/6 

X 

Bay Bush Point 

112/55 

8/< 1 

0 

18/8 

X 

Bluff Point 

54/40 

2 

0 

20/25 

X 

Oldfield 

12/8 

2/< 1 

0 

52/40 

X 

Oldfield 

104/70 

2 

0 

10/4 

X 

Oldfield 

128/60 

12/< 1 

0 

14/6 

X 

Oldfield 

58/40 

8/< 1 

0 

20/23 

X 

Oldfield 

58/40 

38/10 

0 

46/12 

0 

Oldfield 

16/5 

4 

0 

60/40 

X 

Willow Bottom 

0 

0 

0 

122/85 

X 

Nichols 

0 

0 

0 

126/75 

X 

Holton Point 

0 

0 

0 

82/55 

X 

Ebb Point 

2 

0 

0 

64/45 

X 

Spaniard Point 

0 

2 

0 

142/45 

X 

Spaniard Point 

0 

0 

0 

154/80 

X 

Cliff 

0 

2 

0 

130/50 

X 

Cliff 

0 

0 

0 

112/45 

x '[■ - 


(continued) 


106 





1973. Continuation of ltr. from G. E. Krantz of June 8, 1978. 


Bar Markets* Smalls* Spat* Boxes* Associated 

Organisms 
Present t 


X 

X 

X 


Number/Percentage of 1 oyster bushel. 

0 = no organisms detected; X = organisms present. 


Sheep 

0 

0 

0 

26/20 

Mummy Cove 

0 

0 

0 

50/25 

Shippen Creek 

0 

0 

0 

96/25 


107 







1974. Continuation of ltr. from G. E. Krantz of June 8, 1978. 


Bar 

Markets* 

Smalls* 

Spat* 

Boxes* 

Associated 
Organisms 
Present + 

Buoy Rock 

184/80 

102/16 

0 

2 

0 

Blunt 

134/55 

4/1 

0 

12/3 

X 

Hail Point 

90/40 

6 

0 

20/6 

X 

Carpenters Island 

80/30 

6/1 

0 

14/2 

X 

Durdin 

108/53 

12/1 

0 

26 

X 

Horserace 

72/40 

4/< 1 

0 

10/10 

X 

Piney Pt. (Lower) 

90/40 

38/8 

0 

22/5 

X 

Piney Pt. (Upper) 

84/30 

10/1 

0 

46/15 

X 

Bay Bush Point 

56/20 

72/15 

0 

22/5 

X 

Hells Delight 

10/5 

0 

0 

16/10 

X 

Bluff Point 

54/40 

2 

0 

20/25 

X 

Oldfield 

0 

0 

0 

68/25 

X 

Willow Bottom 

0 

0 

0 

74/25 

X 

Nichols 

0 

0 

0 

30/20 

X 

Sand Thistle 

0 

0 

0 

60/15 

X 

Boat House 

0 

0 

0 

88/60 

X 

Drum Point 

0 

0 

0 

28/12 

X 

Holton Point 

0 

0 

0 

82/55 

X 

Ebb Point 

0 

0 

0 

68/25 

X 

Spaniard Point 

0 

0 

0 

132/50 

X 

Cliff 

0 

0 

0 

60/25 

X 

Emory Hollow 

0 

0 

0 

0 

X 


(continued) 







1974. Continuation of ltr. from G. E. Krantz of June 8, 1978. 


Bar 

Markets* 

Smalls* 

Spat* 

* Boxes* 

Associated 
Organisms 
Present t 

Emory Hollow 

0 

0 

0 

0 

X 

Mummys Cove 

2 

0 

0 

30 

0 

Mummys Cove 

0 

0 

0 

42/25 

X 

Shippen Creek 

2 

0 

0 

22/10 

X 

Shippen Creek 

0 

0 

0 

56/40 

X 

* 

Number/Percentage of one 

oyster bushel. 




0 = no organisms detected; X - organisms present. 


109 






1975. Continuation of ltr. from G. E. Krantz of June 8, 1978. 



Bar 


Markets* 

Smalls* 

Spat* 

Boxes* 

Associated 
Organisms 
Present + 

Strong Bay 


162/60 

4/< 1 

0 

8/4 

X 

Ferry 


170/80 

6/< 1 

0 

16/8 

X 

Ferry 


84/15 

0 

0 

20/6 

X 

Ferry 


184/90 

6 

0 

10/6 

X 

Side Shoal 


196/90 

12/4 

0 

8/4 

0 

Buoy Rock 


150/65 

70/12 

0 

2/< 1 

X 

Blunt 


204/85 

0 

0 

16/10 

X 

Blunt 


128/50 

4/1 

0 

20/12 

X 

Blunt 


108/55 

4/< 1 

0 

12/6 

X 

Hail Point 


188/80 

14/2 

0 

8/4 

X 

North of Hail Pt. 

148/45 

16/1 

0 

10/8 

X 

Carpenters 

Island 

122/45 

20/5 

0 

30/12 

X 

Carpenters 

Island 

160/50 

22/4 

0 

10 

X 

Carpenters 

Island 

118/75 

4/1 

0 

4/2 

X 

Carpenters 

Island 

140/45 

54/11 

0 

14/12 

X 

Carpenters 

Island 

238/50 

21/6 

0 

0 

X 

Horserace 


138/60 

24/3 

0 

6/3 

X 

Horserace 


138/50 

60/12 

0 

19 

X 

Piney Point 

70/25 

102/18 

0 

8 

X 

Piney Point 

118/50 

54/10 

0 

30/25 

X 

Bay Bush Point 

8 

52 

0 

14 

0 

Oldfield 


0 

0 

0 

5 

0 

Oldfield 


0 

0 

0 

15 



(continued) 


110 







1975. Continuation of ltr. from G. E. Krantz of June 8, 1978. 


Bar 

Markets* 

Smalls* 

Spat* 

Boxes* 

Associated 
Organisms 
Present t 

Oldfield 

0 

0 

0 

15 

X 

Willow Bottom 

0 

0 

0 

3 

X 

Nichols 

0 

0 

0 

10 

X 

Drum Point 

0 

0 

0 

5 

0 

Holton Point 

0 

0 

0 

48/20 

X 

Spaniard Point 

0 

0 

0 

94/22 

X 

Cliff 

0 

0 

0 

18/6 

0 


Number/Percentage of 1 oyster bushel. 

0 = indicates no organisms detected; X = organisms present. 


Ill 






APPENDIX B 


ACCOUNT OF INTERLABORATORY TESTS ON SPLIT SAMPLES 

This is a chronological account of interactions with Environ¬ 
mental Monitoring and Support Laboratory (EMSL), Office of Re¬ 
search and Development, U.S. Environmental Protection Agency, 
Cincinnati, Ohio. Contact: William L. Budde. 

October 1978 


Two dried sediment samples were each split and respective 
subsamples were sent to EMSL in Cincinnati. They were labeled: 

(1) Chester River, mouth, top sediment; (2) Tenneco Pond sediment 
(>100 ppm dioctyl phthalate). The EMSL facility was selected due 
to its fine reputation in the scientific community. 

January 1979 


EMSL advised this laboratory of their results on the two 
samples: (1) 61 ppm of DEHP in the Chester River, mouth, and 
(2) 1700 ppm DEHP in the Tenneco Pond. No error values nor blank 
values were given. In a telephone conversation prior to this re¬ 
port, EMSL communicated a preliminary verbal report indicating 
6 ppm of DEHP in the Chester River, mouth sediment. (After dis¬ 
covering a computational error, the corrected value was described 
as 58 ppm.) The value this laboratory had determined—prior to 
receiving the EMSL report—was 0.6 ppm DEHP in Chester River, 
mouth sediment and 1200 + 100 ppm DEHP in Tenneco Pond sediment. 

Due to the large discrepancy (2 orders of magnitude) this 
laboratory undertook an extensive battery of experiments to ex¬ 
plore previously undetected systematic errors, and to find a 
possible reason for its low value for DEHP in Chester River, 
mouth sediment. After 2 months of experimentation, we continued 
to obtain values less than 1 ppm. In fact, improvement of our 
methodology produced a value lower than determined previously 
0.1 ppm. 

April 1979 

Agreement for retrial was secured and a second set of split 
samples was sent to EMSL Cincinnati. This time three samples 
were sent: (1) Dry Chester River mouth sediment, (2) Wet Chester 
River mouth sediment, which EPA dried themselves, and (3) "or¬ 
ganic free" clay which we had carefully spiked with DEHP. 


112 





June 1979 


EMSL returned the second set of results and reported a 
drastically lowered result for Chester River, mouth. 


(1) Chester River, mouth 

dried by U. of Maryland 


EPA-Cincinnati 

0.3 mg/kg 
DEHP 


Univ. Maryland 
0.097+0.03 ppm 


DEHP 


(2) Chester River, mouth 
dried by EPA 


0.4 mg/kg 
DEHP 


(3) Attapulgite, spiked 


121 mg/kg 
DEHP 


200+10 ppm,DEHP 


In a telephone conversation prior to this report, EMSL commented 
that during the time of the first exchange, EPA Cincinnati had 
been experiencing some contamination problems with their homo- 
genizer used in sediment extraction. This is a probable explana¬ 
tion for the excessively high values found by EMSL in the origin¬ 
al exchange. In the report of June 1979, signed by William 
Budde, it was mentioned that EMSL procedures were the same as 
before, except that they had used a new Tissumizer (R) by Tekmar. 

While there is still some difference between the values de¬ 
termined by each laboratory in this study, we believe they are 
within acceptable limits especially considering the history of 
trace organic interlaboratory comparisons (Hilpert et al., 1978, 
Hertz et al., 1979). 


Hertz, H. S., L. R. Hilpert, W. E. May, S. A. Wise, J. M Brown, 

S. N. Chesler, and F. R. Guenther. Special Technical Publication 
686, American Society for Testing and Materials, 1979. 

Hilpert, L. R., W. E. May, S. A. Wise, S. N. Chesler, and H. S. 
Hertz. Interlaboratory comparison of determinations of trace 
level petroleum hydrocarbons in marine sediment. Anal. Chem. 
50:458-463, 1978. 


113 




APPENDIX C 


METHODS FOR WATER QUALITY MEASUREMENTS IN TIN STUDY 

Water temperature was measured with the temperature probe on 
the YSI oxygen meter used to determine dissolved oxygen. 
Measurement of pH was with a Coleman meter, and salinity was 
measured using a Wheatstone conductivity bridge. 


114 




























































































































232 91 



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BINDERY INC. 


9t MAY 91 



N. MANCHESTER, 
INDIANA 46962 


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